Role of nitric oxide on ATP-induced Ca2+ signaling in outer hair cells of the guinea pig cochlea

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a v a i l a b l e a t w w w. s c i e n c e d i r e c t . c o m

w w w. e l s e v i e r. c o m / l o c a t e / b r a i n r e s

Research Report

Role of nitric oxide on ATP-induced Ca 2+ signaling in outer hair cells of the guinea pig cochlea Jing Shen a , Narinobu Harada a,b,⁎, Hiroko Nakazawa a , Toshishiko Kaneko a , Masahiko Izumikawa a , Toshio Yamashita a a

Hearing Research Laboratory, Department of Otolaryngology, Kansai Medical University, Fumizonocho 10-15, Moriguchi, Osaka 570-8507, Japan b Harada Ear Institute, Tomoi 2-34-27, Higashiosaka, Osaka 577-0816, Japan

A R T I C LE I N FO

AB S T R A C T

Article history:

Recently, a negative feedback effect of nitric oxide (NO) on the adenosine 5′-triphosphate

Accepted 27 December 2005

(ATP)-induced Ca2+ response has been described in cochlear inner hair cells. We here

Available online 28 February 2006

investigated the role of NO on the ATP-induced Ca2+ response in outer hair cells (OHCs) of the guinea pig cochlea using the NO-sensitive dye DAF-2 and Ca2+-sensitive dye fura-2. G

Keywords:

Extracellular ATP induced NO production in OHCs, which was inhibited by

Outer hair cell

nitroarginine methyl ester (L-NAME), a non-specific NO synthase (NOS) inhibitor, and

Nitric oxide

suramin, a P2 receptor antagonist. ATP failed to induce NO production in the Ca2+-free

ATP

solution. S-nitroso-N-acetylpenicillamine (SNAP), a NO donor, enhanced the ATP-induced

L-N

-

Cochlea

increase of the intracellular Ca2+ concentrations ([Ca2+]i), while L-NAME inhibited it. SNAP

Intracellular Ca2+

accelerated ATP-induced Mn2+ quenching in fura-2 fluorescence, while L-NAME suppressed it. 8-Bromoguanosine-cGMP, a membrane permeable analog of cGMP, mimicked the effects of SNAP. 1H-[1,2,4]oxadiazole[4,3-a] quinoxalin-1-one, an inhibitor of guanylate cyclase and KT5823, an inhibitor of cGMP-dependent protein kinase inhibited the ATP-induced [Ca2+]i increase. Selective neuronal NOS inhibitors, namely either 7-nitro-indazole or 1-(2trifluoromethylphenyl) imidazole, mimicked the effects of L-NAME regarding both ATPinduced Ca2+ response and NO production. Immunofluorescent staining of neuronal nitric oxide synthase (nNOS) in isolated OHCs showed the localization of nNOS in the apical region of OHCs. These results suggest that the ATP-induced Ca2+ influx via a direct action of P2X receptors may be the principal source for nNOS activity in the apical region of OHCs. Thereafter, NO can be produced while conversely enhancing the Ca2+ influx via the NOcGMP-PKG pathway by a feedback mechanism. © 2006 Elsevier B.V. All rights reserved.

1.

Introduction

Nitric oxide (NO), a gaseous membrane permeate messenger, has been thought to play an important role in signaling

processing as a neurotransmitter or a neuromodulator in the olfactory system (Collmann et al., 2004), visual system (Wang et al., 2003), and central neural system (Esplugues, 2002). NO is synthesized by NO synthase (NOS). In the peripheral auditory

⁎ Corresponding author. Hearing Research Laboratory, Department of Otolaryngology, Kansai Medical University, Fumizonocho 10-15, Moriguchi, Osaka 570-8507, Japan. Fax: +81 6 6726 8111. E-mail address: [email protected] (N. Harada). 0006-8993/$ – see front matter © 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.brainres.2005.12.129

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organ, Ca2+-dependent constitutive NOS, neural NOS (nNOS), and endothelial NOS (eNOS) have been demonstrated by means of immunocytochemistry (Franz et al., 1996; Gosepath et al., 1997; Heinrich et al., 2004). Those studies showed that nNOS and eNOS were localized in several types of cells in the cochlea, such as inner hair cells (IHCs), outer hair cells (OHCs), spiral ganglion neurons (SGNs), and supporting cells (Franz et al., 1996; Gosepath et al., 1997; Heinrich et al., 2004). Recent studies using the NO-sensitive dye DAF-2 showed NO production to occur in several types of cells in the guinea pig cochlea (Shen et al., 2003; Shi et al., 2001; Takumida and Anniko, 2001; Yukawa et al., 2005). These results suggest that NO may play an important role in the inner ear (Takumida and Anniko, 2002). IHCs and OHCs are the two sensory receptors of the mammalian cochlea and exhibit innervation and functional properties. OHCs are innervated by large myelinated efferents and are exhibit somatic motility (Dallos, 1992; He et al., 2004) that is thought to contribute to the enhancement of cochlear frequency selectivity by generating negative damping (Dallos and Corey, 1991), although recent work also suggests that the hair bundle generates forces that could be involved in this process (Robles and Ruggero, 2001). IHCs appear to perform a more passive role and are innervated by the majority of afferent auditory nerve fibers, hence giving rise to the major neural output of the cochlea. Previous studies also suggested that the elevation of intracellular Ca2+ concentrations ([Ca2+]i) affects OHC motility (Frolenkov et al., 2003; Szonyi et al., 2001). It has been suggested that the effects of NO are involved in the regulation of [Ca2+]i through the NO-cGMP-PKG signaling pathway in the sensory system of olfaction (Breer and Shepherd, 1993) and vision (Cudeiro and Rivadulla, 1999). Morphological studies have shown this pathway to also exist in the cochlea (Heinrich et al., 2000; Takumida et al., 2000). However, little is known about the role of the NO-cGMP-PKG pathway in auditory neurotransmission. Extracellular adenosine 5′-triphosphate (ATP) has been described as a neurotransmitter or a neuromodulator in auditory neurotransmission (Housley et al., 2002). ATP has been reported to induce an increase of [Ca2+]i in OHCs (Ashmore and Ohmori, 1990; Mammano et al., 1999), IHCs (Shen et al., 2003, 2005; Sugasawa et al., 1996; Yamashita et al., 1993), and SGNs (Cho et al., 1997; Yukawa et al., 2005). These previous studies suggested that ATP may affect auditory neurotransmission by modulating [Ca2+]i in the cochlea. Previous studies have also shown OHCs to possess both ligand-gated ionotropic P2X receptors and G-protein-coupled metabotropic P2Y receptors (Ashmore and Ohmori, 1990; Mammano et al., 1999; Raybould and Housley, 1997; Van den Abbeele et al., 1996). We have recently reported that NO inhibits the ATP-induced Ca2+ response via the NO-cGMP-PKG pathway in IHCs by a negative feedback mechanism, while NO enhances the ATPinduced Ca2+ response in SGNs by a positive feedback mechanism (Shen et al., 2003, 2005; Yukawa et al., 2005). Our data demonstrated that a cross-talk between NO and ATP may exist in the afferent auditory signal transduction. We also suggested that NO may play a crucial role in the process of auditory afferent signaling transduction in the cochlea. However, the role of NO in cellular signaling transduction of OHCs has not yet been elucidated. We therefore investigated whether ATP can induce NO production, which affects ATP-induced Ca2+ signaling in

OHCs. We also investigated the role of the NO-cGMP-PKG pathway on the ATP-induced Ca2+ signaling in OHCs.

2.

Results

2.1.

ATP-induced endogenous NO production in OHCs

The application of 100 μM ATP caused a gradual increase in DAF-2 fluorescence, which reached a plateau at about 86% above baseline after 10–12 min in the continued presence of extracellular ATP (Figs. 1A, B). The average DAF-2 ratio (F/F0) at a maximal level was about 1.86 ± 0.16 (n = 12). A non-specific NO synthase inhibitor, 200 μM L-NAME (Scrogin et al., 1998) inhibited the ATP-induced increase in DAF-2 fluorescence to 25 ± 4% of ATP alone (P b 0.01, 1.22 ± 0.05 vs. 1.86 ± 0.16; n = 13; Figs. 1B, C), while an inactive analog D-NAME (200 μM) did not inhibit it (P N 0.05, 1.95 ± 0.06 vs. 1.86 ± 0.16; n = 10; Fig. 1C). PSS alone did not induce any apparent changes in DAF-2 fluorescence in the presence of 100 μM L-arginine (F/F0: 1.03 ± 0.04; n = 8; Figs. 1B, C). Furthermore, 100 μM suramin, an antagonist of P2-purinergic receptors, inhibited the ATPinduced increase of DAF-2 fluorescence in OHCs to 32 ± 7% of ATP alone (P b 0.01, 1.28 ± 0.17 vs. 1.86 ± 0.16; n = 10; Figs. 1B, C). To test whether an increase in [Ca2+]i is required for ATPinduced NO production, we performed the experiment in Ca2+free solution. In the absence of extracellular Ca2+, ATP failed to induce a significant increase in DAF-2 fluorescence compared to PSS alone (P N 0.05, 1.04 ± 0.05 vs. 1.03 ± 0.04; n = 10; Figs. 1B, C). The calmodulin antagonist, calmidazolium abolished the ATP-induced NO production in OHCs (P b 0.01, 1.02 ± 0.04 vs. 1.86 ± 0.16; n = 7; Figs. 1C, D).

2.2. Simultaneous measurement of the ATP-induced NO production and [Ca2+]i changes In order to investigate whether ATP-induced NO production is involved the elevation of [Ca2+]i, simultaneous measurement of the ATP-induced NO production and [Ca2+]i changes were performed in an isolated OHC loaded with both DAF-2 DA and fura-2/AM. The temporal relationship between the ATPinduced NO production and increase in [Ca2+]i was simultaneously investigated in seven cells of isolated OHCs. Both NO production and the increase in [Ca2+]i induced by ATP were observed in OHCs (Fig. 2). 100 μM ATP caused a rapid transient increase in [Ca2+]i of OHCs, while the DAF-2 fluorescence increased more slowly than the [Ca2+]i increase and lagged behind the Ca2+ response (Figs. 2A, B).

2.3. NO enhances the ATP-induced [Ca2+]i increase by a feedback mechanism To investigate whether NO can modulate the ATP-induced Ca2+ response in OHCs, we investigated the effects of a NO donor, SNAP (Garcia-Pascual et al., 1999) and L-NAME on the [Ca2+]i changes induced by ATP. The application of extracellular ATP (100 μM) increased [Ca2+]i in OHCs up to peak values of 523 ± 172 nM from a resting level of 108 ± 35 nM (n = 23). The response to a subsequent application ATP after a 15-min interval was similar to the amplitude of the first response in OHCs as shown

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previously (Shen et al., 2005) (Fig. 3D). The amplitude of the second [Ca2+]i increase was 98.2 ± 0.09% of the first response (P N 0.05, second response vs. first response, n = 10; paired Student's t test. Figs. 3C, I). Therefore, two applications of ATP at intervals of at least 15 min were used in the present study. The ATP-induced Ca2+ responses were therefore reversible and repeatable in the present study. In isolated cochlear OHCs, 100 μM SNAP enhanced the ATP-induced [Ca2+]i increase to 184 ± 28% of the first response (P b 0.01 vs. first response; n = 12; paired Student's t test. Figs. 3E, I), while 200 μM L-NAME inhibited it to 58 ± 12% of the first response (P b 0.01 vs. first response; n = 9; paired Student's t test. Figs. 3G, I). In contrast to OHCs, 100 μM SNAP inhibited the ATP-induced [Ca2+]i increase to 69 ± 14% of the first response in IHCs (P b 0.01 vs. first response, n = 8; paired Student's t test. Figs. 3F, J), while 200 μM 2+ L -NAME enhanced the ATP-induced [Ca ] i increase to 203 ± 50% of the first response in IHCs (P b 0.01 vs. first response, n = 15; paired Student's t test. Figs. 3H, J). Bath application of ATP caused a slow increase in DAF-2 fluorescence with a lag time compared with fura-2 fluorescence (Fig. 2). Therefore, this recording protocol may not be able to detect the rapid changes in endogenous NO production. To confirm whether ATP can rapidly induce NO production, we locally applied ATP to OHCs using a puffing ejection technique. However, in 8 cells, local application of ATP evoked an initial increase in DAF-2 fluorescence within 2–6 s (4.0 ± 1.6 s) after stimulation. Fig. 4 shows a representative rapid increase in DAF-2 fluorescence induced by the local application of ATP. These results indicate that ATP can rapidly induce NO production, which thereby enhances the Ca2+ response. We also investigated whether SNAP alone can induce NO production. 100 μM SNAP alone enhanced the DAF-2 fluorescence (Fig. 5A), while it did not elicit any detectable changes in [Ca2+]i in OHCs (Fig. 5B). SNAP can thus be used as a stimulator for the augmentation of ATP-induced NO production.

2.4.

Fig. 1 – ATP-induced NO production in cochlear OHCs. (A) A bright field image of single OHC (left). Fluorescence images of NO production induced by 100 μM ATP at prestimulation and 15 min after ATP stimulation (middle and right). Scale bar, 10 μm. The calibration bar on the right shows the DAF-2 fluorescence intensity in pseudocolor. (B) Typical time course of the normalized changes in DAF-2 fluorescence induced by extracellular ATP (100 μM) in the OHCs with several treatments. Control perfusion with PSS alone did not induce any significant increase in DAF-2 fluorescence in OHCs. Both 200 μM L-NAME and 100 μM suramin inhibited the ATP-induced NO production. ATP failed to induce NO production in the presence of calmodulin inhibitor calmidazolium (10 μM) or in the absence of extracellular Ca2+. The closed horizontal bar at the bottom of trace indicates the timing of ATP application. (C) Histogram summarizing the normalized average maximum in panel B. 100 μM D-NAME, inactive isomer of L-NAME did not show any effects on the ATP-induced NO production. The number of the cells tested is shown in parenthesis. Error bar shows SD. *P b 0.01 vs. ATP-induced NO production as a control (one-way ANOVA).

NO enhances the ATP-induced Mn2+ influx

To investigate whether NO enhances the ATP-induced Ca2+ influx by a feedback mechanism, experiments were performed using Mn2+ quenching of fura-2 fluorescence. As shown in Fig. 6, application of 50 μM Mn2+ caused a decline of fura-2 fluorescence in OHCs in the presence of ATP (n = 9) compared to control measurements with PSS. This indicates that ATP triggered Mn2+ influx through the ATP-gated channels. In the presence of 100 μM SNAP, the ATP-induced Mn2+ quenching rate of fura-2 fluorescence in OHCs was significantly increased to 345 ± 68% of ATP alone (P b 0.001 vs. ATP alone, n = 9; Fig. 6). Pretreatment of the OHCs with 200 μM L-NAME significantly suppressed the ATP-induced Mn2+ quenching rate to 20 ± 3% of ATP alone (P b 0.001 vs. ATP alone, n = 8. Fig. 6).

2.5. Involvement of the NO-cGMP-PKG pathway in the NO-mediated feedback effect of the ATP-induced Ca2+ response To investigate whether the NO-cGMP-PKG signaling pathway is involved in the effect of NO on the ATP-induced Ca2+ response, a membrane permeable cGMP, 8-Br-cGMP, and a selective inhibitor of the NO-sensitive guanylate cyclase, ODQ (Boulton et al., 1995) and an inhibitor of cGMP-dependent protein

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Fig. 2 – Simultaneous measurement of NO production and [Ca2+]i changes in OHCs. (A) Pseudocolor displays of the ATP-induced [Ca2+]i increase (top panel) in parallel with the ATP-induced NO production (bottom panel) in one OHC. The fura-2 and DAF-2 fluorescence signals were alternately excited at 340 nm and 480–490 nm, respectively. The fluorescence images of DAF-2 and fura-2 were taken at the times indicated above each pair of images. Note that the [Ca2+]i increase was observed 20 s after the onset of 100 μM ATP stimulation, whereas no significant change in DAF-2 fluorescence was observed in this cell (b). A steep increase in DAF-2 fluorescence was observed 240 s after ATP stimulation, while the fura-2 fluorescence declined to basal levels (d). Scale bar is 10 μm. (B) Time course changes of NO production and [Ca2+]i increase in panel A. The DAF-2 fluorescence intensities are normalized relative to the initial value (F/F0; right vertical axis) and are plotted in red; changes in fura-2 fluorescence intensity are expressed by the ratio of F340/F360 (left vertical axis) and are plotted in black. The horizontal bar at the bottom of trace indicates the timing of ATP application.

kinase (PKG), KT5823 (Burkhardt et al., 2000) were used in the following experiments. 100 μM 8-Br-cGMP enhanced the amplitude of ATP-induced [Ca2+]i increase to 149 ± 33% of the control (P b 0.001 vs. first response to ATP, n = 16; paired Student's t test; Figs. 7A, D). 10 μM ODQ and 1 μM KT5823 inhibited the amplitude of ATP-induced [Ca2+]i increase to 60 ± 17% and 64 ± 18% of the control, respectively (ODQ: P b 0.001 vs. first response to ATP, n = 10; KT5823: P b 0.001 vs. first response to ATP, n = 12. Paired Student's t test; Figs. 7B, C, D).

2.6.

inhibited the amplitude of ATP-induced [Ca2+]i increase to 54 ± 9% and 58 ± 11% of the first response to ATP, respectively (7-NI: P b 0.001, normalized ratio: 0.54 ± 0.09 vs. 0.98 ± 0.09, n = 14; TRIM: P b 0.001, 0.58 ± 0.11 vs. 0.98 ± 0.09, n = 13; Fig. 8B). There was no significant difference between the effect of 7-NI and L-NAME ([NO]: P N 0.05, 1.23 ± 0.06 vs. 1.22 ± 0.05; [Ca2+]i: P N 0.05, 0.54 ± 0.09 vs. 0.58 ± 0.12), TRIM and L-NAME ([NO]: P N 0.05, 1.25 ± 0.07 vs. 1.22 ± 0.05; [Ca2+]i: P N 0.05, 0.58 ± 0.11 vs. 0.58 ± 0.12) on both the ATP-induced NO production and [Ca2+]i increase (Figs. 8A, B).

ATP-induced NO production is derived from nNOS

To clarify the contribution of NOS isoforms to the ATP-induced NO production in OHCs, we compared the effects of L-NAME, a non-selective NOS inhibitor, with the two other relatively selective inhibitors for nNOS, 7-NI (Ayajiki et al., 2001), and TRIM (Handy and Moore, 1997) on both ATP-induced NO production and [Ca2+]i increase. 100 μM 7-NI and 100 μM TRIM inhibited the amplitude of ATP-induced NO production to 26 ± 5% and 29 ± 6%of ATP alone, respectively (7-NI: P b 0.001, 1.23 ± 0.06 vs. 1.86 ± 0.16, n = 15; TRIM: P b 0.001, 1.25 ± 0.07 vs. 1.86 ± 0.16, n = 16; Fig. 8A). 100 μM 7-NI and 100 μM TRIM also

2.7. Interaction between nNOS and P2X2 receptor on the ATP-induced NO production To investigate localization of nNOS and P2X receptors in OHCs, we performed co-localization studies by means of immunofluorescent staining for nNOS and double staining for nNOS/ P2X2 receptors in isolated OHCs. As shown in Fig. 9A, intense immunofluorescent staining for nNOS was observed in the apical region of OHCs including the stereocilia and the cuticular plate and infracuticullar network. No apparent fluorescent labeling

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Fig. 3 – Effect of NO on the ATP-induced [Ca2+]i increase in OHCs and IHCs. (A, B) DIC images of single OHC (A) and IHC (B) from the guinea pig cochlea. (C, D) A representative recordings of the [Ca2+]i changes induced by sequential 100 μM ATP application in OHCs and IHCs. There is no difference in the magnitude of [Ca2+]i evoked by two consecutive ATP applications with a 15-min interval. (E–H) Effect of SNAP and L-NAME on the ATP-induced [Ca2+]i increase in OHCs and IHCs. Pretreatment of the OHCs with 100 μM SNAP enhanced the ATP-induced [Ca2+]i increase (E), while 100 μM SNAP inhibited it in IHCs (F). 200 μM L-NAME inhibited the ATP-induced [Ca2+]i increase in OHCs (G), while 200 μM L-NAME enhanced the ATP-induced [Ca2+]i increase in IHCs (H). (I, J) Summary histogram of the effects of NO on ATP-induced [Ca2+]i increase in panels C–H. Values of the second Ca2+ response to ATP plus agent are normalized to the control (first) response. The numbers of cells tested are given in parentheses. Error bar shows SD. *P b 0.01 vs. ATP-induced Ca2+ response as a control (one-way ANOVA). The horizontal bar at in each trace indicates the timing of drug application.

could be detected in the negative control, which was performed by either omitting the primary nNOS antibody (Fig. 9A) or incubating the cells with anti-nNOS antibody blocking peptide (Fig. 9A). As for nNOS/P2X2 double staining, P2X2 immunofluorescence labeling was mainly restricted to the apical region of OHCs (Fig. 9B), where nNOS was intensely stained (Fig. 9B). These results directly provided morphological evidence, suggesting the possible interaction between nNOS and P2X2 receptor in OHCs. To confirm the specificity of the nNOS and P2X2, antibodies used in immunocytochemistry, Western blot analysis was performed. Immunoblots probed with the antibodies against nNOS and P2X2 revealed a ∼155-kDa and a ∼64-kDa band in the organ of Corti, respectively. These results indicate that nNOS and P2X2, antibodies used in the present study, are specificity for nNOS and P2X2, respectively (Fig. 9C).

2.8.

Localization of ATP-induced NO production in OHCs

Because of the co-localization of nNOS and P2X2 in the apical region of OHCs, we investigated whether this region plays a special role in the ATP-induced endogenous NO production. In the present study, NO production induced by ATP was frequently observed in the apical region of OHCs as shown in Fig. 1. However, the thickness of isolated OHCs coupled with observation by epifluorescence rather than confocal microscopy may have resulted in the obscuration of the apical signal, especially as cells failed to show the apical changes associated with NO production. Therefore, we analyzed the spatial and temporal distribution of changes in NO evoked by local application of ATP using real-time confocal microscope. The local application of 100 μM ATP to the apical region of the OHCs induced a notable increase in DAF-2 fluorescence in the OHCs (Fig. 10A; the upper trace). The application of 100 μM ATP to

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Fig. 4 – NO production induced by local application of ATP. 100 μM ATP was locally applied to the apical region of the OHCs by puff pipettes (tip diameter of 2–5 μm) held about 10–20 μm away from the OHCs. Note that the NO production was apparent at 2–4 s after local application of extracellular ATP. Local application of PSS did not induce NO production as a control. the middle region of the OHCs only induced only a slight increase in DAF-2 fluorescence (Fig. 10A; the middle trace). There were no significant increases in DAF-2 fluorescence in the OHCs when 100 μM ATP was applied to the basal region of the OHCs (Fig. 10A; the lower trace). The DAF-2 fluorescence intensities were increased initially in the apical region of the OHCs during ATP stimulation using confocal microscopy (Fig. 10B).

3.

Discussion

The present study suggests that NO enhances the ATP-induced Ca2+ influx via the NO-cGMP-PKG pathway in OHCs by a positive feedback mechanism. Our results also suggest that the ATP-induced Ca2+ influx can be the principal source for nNOS activity which co-localizes with ionotropic P2X receptors in the apical region of OHCs. To our knowledge, this is the first study to demonstrate that nNOS coupled to P2X receptor in cochlear OHCs plays a role in regulating the Ca2+ signaling in OHCs. We previously reported that ATP can induce NO production in cochlear IHCs, which in turn inhibits the ATP-induced [Ca2+]i increase by a negative feedback mechanism (Shen et al., 2003,

2005). We suggested that NO may play an important role in the afferent modulation of the cochlea. Our present study also suggests that NO plays a role not only in afferent modulation but also in efferent modulation of the cochlea since the medial efferent system originates in the medial superior olivary complex and innervates OHCs (Dallos, 1992). Previous study suggests that DAF-2 fluorescence may be influenced by the changes of Ca2+ concentrations, which is over 2 mM Ca2+ (Broillet et al., 2001). In the present study, the ATP-induced [Ca2+]i increase never exceeded micromolar levels in OHCs. ATP-induced NO production was abolished in the absence of extracellular Ca2+, while ATP could induce [Ca2+]i increase by release from internal stores (Mammano et al., 1999). If the increase in DAF-2 fluorescence was affected by the [Ca2+]i increase (Broillet et al., 2001), the increase of DAF-2 fluorescence would be accompanied with the ATP-induced [Ca2+]i increase in OHCs. Furthermore, a recent study showed that the sensitivity of DAF-2 to NO gas was independent of cytoplasmic Ca2+ and Mg2+ at physiological concentrations (Suzuki et al., 2002). Therefore, the influence of those factors on the DAF-2 fluorescent signal could be ruled out in the present study. Our results also suggest that NO production may be due to an increase in the level of [Ca2+]i by the Ca2+ influx through the activation of P2X receptors. The simultaneous measurement of NO production and [Ca2+]i changes provided evidence that the elevation of [Ca2+]i may be responsible for the NOS activity in OHCs. Our findings are consistent with the notion that the elevation of [Ca2+]i was necessary for NO production (Dedkova and Blatter, 2002; Li et al., 2003; Lin et al., 2000). The ATP-induced NO production has been reported in aortic endothelial cells (Dedkova and Blatter, 2002; Kimura et al., 2004), astrocytes (Li et al., 2003), and cochlear IHCs (Shen et al., 2003), SGNs (Yukawa et al., 2005). In the cochlea, both ATP and NO have been considered to be candidates for neurotransmitters or neuromodulators, and they have therefore been implicated in the signaling processing of auditory system (Sagami et al., 2001; Housley et al., 2002; Kennedy et al., 2005). Interestingly, we report here an interaction between ATP and NO in cochlear OHCs. Together with our previous studies about the interaction of ATP and NO in cochlear IHCs and SGNs (Shen et al., 2003; Yukawa et al., 2005), we presume a novel regulatory loop for the NO-mediated auditory information processing in the cochlea. Regarding the

Fig. 5 – Effect of SNAP on NO production and [Ca2+]i changes. (A) Application of 100 μM SNAP induced an increase in DAF-2 fluorescence intensity, thus indicating NO production in OHCs. (B) 100 μM SNAP did not induce a [Ca2+]i increase in OHCs.

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Fig. 6 – Effect of NO on the ATP-induced Mn2+ influx. (A) 100 μM SNAP enhanced the rate of Mn2+ quenching while 200μM 2+ L-NAME decreased it. 100 μM ATP was added 4 min prior to application of 50 μM Mn . Experiments were performed in 2+ Ca -free solution. SNAP or L-NAME was added at least 15 min before ATP stimulation. The horizontal bar in each trace in panel A indicates the timing of drug application. (B) Histogram summarizing the effects of L-NAME and SNAP on the ATPinduced Mn2+ influx. Each bar represents the mean ± SD. The number of OHCs tested in separate experiments is indicated in parentheses. *P b 0.01 vs. control (one-way ANOVA). opposite effects of NO on Ca2+ signaling in OHCs and IHCs, they could be relevant to the roles of these two classes of sensory cells for hearing in the cochlea. We cannot rule out the possibility that NO may also enhance the elevation of [Ca2+]i by releasing the Ca2+ from the internal stores. The present study suggests that nNOS, rather than eNOS, may be the main isoform of NOS for the ATP-induced NO production in OHCs. Recently, Heinrich et al. reported that the quantitative immuno-electron microscopic analysis revealed cellular differences in the degree of eNOS and nNOS expression in OHCs (Heinrich et al., 2004). They suggested that these differences might be connected to the different subcellular binding properties of nNOS and eNOS. They concluded that these two constitutive NOS (nNOS and eNOS) might be located at different subcellular sites and might be regulated by the different Ca2+ response. A previous study also showed that NO production induced by glutamate in cultured retinal ganglion neurons was considered to be mainly associated with nNOS and eNOS did not appear to play a major role in NO production because the NO production was inhibited by 7-NI, while immunohistochemically both nNOS and eNOS were expressed in retinal ganglion neurons (Tsumamoto et al., 2002). They speculated that the synthesis of NO by eNOS may represent a compensatory mechanism in the absence of nNOS. Therefore, the different role of eNOS and nNOS may also be associated with different cellular upstream and downstream signaling molecules, which respond to different extracellular signals and are responsible for different cellular functions in OHCs. However, we cannot exclude the possibility that eNOS may also affect the ATP-induced NO production and Ca2+ response in OHCs. Further study will be needed to detect the cellular distribution of eNOS in isolated OHCs. Our findings that nNOS and P2X receptors were co-localized at the apical region of OHCs are consistent with a recent study, which showed the co-localization of nNOS and P2X2 receptors in the neurons of hypothalamus and brain stem of rat (Yao et al., 2003). These findings are also in line with the observations in real-time by confocal laser microscopy that ATP-induced NO production was initially restricted to the apical region. Our

morphological and functional evidence suggest that nNOS interacts with P2X2 receptors. Recent study also suggests that active hair-bundle motion may have an important role in cochlear amplification in mammals (Kennedy et al., 2005). Therefore, we suggest that interaction between nNOS and P2X2 in the apical region of OHCs, including cilia and hair bundle, may participate to the control of amplification by active hair bundle motion. The functional significance and role of nNOS in active hair bundle motion of OHCs should therefore be explored. It has been shown that the apical region of OHCs, including stereocilia and the cuticular plate, is abundant in calmodulin (Furness et al., 2002), P2X receptors (Housley et al., 1999), and plasma membrane Ca2+-ATPase 2a subunit (Dumont et al., 2001). Calmodulin binding to nNOS is reported to be essential for the nNOS activity, by which electron transfer is triggered (Sagami et al., 2001). Our results showed that calmidazolium, a calmodulin antagonist, abolished the ATP-induced NO production. P2X receptors are implicated in the ATP-induced Ca2+ influx, while the plasma membrane Ca2+ ATPase contributes to Ca2+ homeostasis by extruding Ca2+ from the cytoplasm. Our findings thus suggest that these proteins co-localized in the apical region of OHCs might be involved in the upstream and downstream signaling for nNOS, thereby accounting for the activation and functions of nNOS in OHCs. Confocal imaging of NO production in an isolated OHC showed that ATP-induced NO production was detected in the apical region of the OHCs, however, NO production was remarkably observed in the region below the cuticular plate. A previous study suggested that a high expression of NOS does not always imply a high NO production in the hippocampus and cortex during NMDA stimulation (Kojima et al., 1998). Therefore, differences between the ATP-induced NO production and high expression of nNOS in OHCs may be due to the different functional roles of NO as previously suggested (Kojima et al., 1998). Another possible reason might be the occurrence of NO production only in localized regions of the cell as soon as it is stimulated. Such a small amount of DAF-2 production within the cell was not detectable based on the imaging system used in the present study. Further study, using a high-speed imaging system,

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demonstrating that nNOS is activated by the ATP-induced Ca2+ influx via the action of P2X receptors in the apical region of OHCs. In turn, NO was produced and then enhanced the ATP-induced Ca2+ influx via the NO/cGMP/PKG pathway by a positive feedback mechanism. Moreover, our findings also suggest that both ATP and NO, as the neurotransmitter or neuromodulator in the cochlea, induce the co-regulation of Ca2+ homeostasis in OHCs.

Fig. 7 – Augmentation of the ATP-induced [Ca2+]i increase by NO is mediated by the NO/cGMP/PKG pathway. (A) 100 μM 8-Br-cGMP, a membrane-permeant cGMP analogue, enhanced the ATP-induced [Ca2+]i increase. (B) 10 μM 7ODQ, an inhibitor of guanylate cyclase, inhibited the ATP-induced [Ca2+]i increase. (C) 1 μM KT5823, a PKG inhibitor inhibited the ATP-induced [Ca2+]i increase. (D) Summary of the statistical results on the ATP-induced [Ca2+]i changes in panels A, B, and C. Values of the second Ca2+ response to ATP plus agent (8-Br-cGMP, ODQ or KT5823) are normalized to the first response, and second response to ATP alone was normalized to the first response as control. The number of OHCs tested in separate experiments is indicated in parentheses. Error bar shows SD. *P b 0.01 compared to the control. Each inhibitor or analogue was added at least 15 min before second ATP stimulation. The horizontal bar in each trace indicates the timing of drug application.

is thus called for to fully understand the temporal and spatial production of NO in OHCs induced by ATP. In conclusion, the present study made a new discovery of the interaction between nNOS and P2X2 in OHCs while also

4.

Experimental procedures

4.1.

Preparation of outer hair cells

This study was reviewed by the Committee for Ethics for Animal Experiments of Kansai Medical University and was carried out under the Guidelines for Animal Experiments of Kansai Medical University. All experiments also confirmed to international guidelines on ethical use of animals. Outer hair cells (OHCs) were acutely isolated from the guinea pig cochlea without enzymatic treatment as described previously (Harada et al., 1993). Briefly, albino guinea pigs weighting 250–360 g with good Preyer's reflexes were deeply anesthetized and decapitated. The bilateral temporal bones were rapidly removed and placed into physiological standard solution (PSS) containing (mM) NaCl, 150; KCl, 5; MgCl2, 1; dglucose, 5; HEPES, 10; CaCl2, 2, adjusted to a pH 7.4 and 300 mosM by adding distilled water. Then the bullas were immediately opened, and the bony wall of the cochlea was chipped away with a surgical blade. The organ of Corti was further carefully dissected from the modiolus, and then the tissue strips were harvested and gently transferred into a 300 μl experimental chamber coated with Cell-Tak (Cosmo-Bio, Tokyo, Japan) using a micropipette. The cells were mechanically isolated at room temperature using gentle trituration and the isolated OHCs allowed to settle onto the bottom of the chamber for 5–10 min. The viability of OHCs was determined by the following morphological criteria: a uniform cylindrical cell shape, the absence of cytoplasmic Brownian motion, the basal location of the nucleus and undisturbed stereocilia. For some experiments, the IHCs were also isolated as described previously (Shen et al., 2003, 2005).

4.2.

[Ca2+]i measurements

Intracellular Ca2+ concentrations ([Ca2+]i) in OHCs were measured using the HisCA/Argus Imaging System (Hamamatsu Photonics, Hamamatsu, Japan), as described previously (Cho et al., 1997; Shen et al., 2003). In brief, the isolated OHCs were loaded with 2 μM fura-2 acetoxy methyl ester (fura-2/AM) (Dojindo, Kumamoto, Japan) for 30 min at room temperature. Fura-2 loaded OHCs were alternately illuminated by 340-nm and 380-nm excitation. The emitted light was monitored after passage through a 510-nm cut-off filter (a 20-nm band-pass). The fluorescence ratio (340 nm/380 nm) was calculated and converted into absolute values of [Ca2+]i based on the formula described by Grynkiewicz et al. (1985), ½Ca2þ i ¼ Kd ½ðR  Rmin Þ=ðRmax  RÞF0 =Fs Rmin, Rmax, Kd, F0/Fs values were determined to be 0.22, 6.57, 224 nM, and 8.8, using ethylene glycol-bis (β-aminoethyl Ether) N,

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Fig. 8 – Effect of L-NAME, 7-NI and TRIM on the ATP-induced NO production and [Ca2+]i changes. (A, B) Histogram summarizing the comparative effects of 7-NI and TRIM, selective inhibitors of for nNOS, and L-NAME a non-selective NOS inhibitor on the ATP-induced NO production (A) and the ATP-induced [Ca2+]i increase (B). No significant differences were observed between the effects of 7-NI, TRIM, and L-NAME on both the ATP-induced NO production and Ca2+ response. The numbers of cells tested are given in parentheses. Error bar shows SD. *P b 0.01 vs. control. N.S. indicates not significant (one-way ANOVA).

N,N′,N′-tetraacetic acid-Ca2+ buffer solution with known concentrations of Ca2+ containing 25 μM fura-2 pentapotassium salt. Measurements of [Ca2+]i in IHCs were also carried out as described above.

4.3.

Measurements of intracellular NO production

The protocol for measuring the NO production was similar to that we reported elsewhere (Shen et al., 2003; Yukawa et al., 2005). The cells were loaded with 5 μM 4,5-diaminofluorescein diacetate (DAF-2DA, Daiichi Pure Chemicals Co., Ltd., Tokyo, Japan) for 30 min at room temperature. The DAF-2 fluorescence in the OHCs was imaged using an inverted fluorescent

microscope (TMD-2, Nikon, Tokyo, Japan) equipped with a xenon lamp, excitation (485–495 nm) and emission (510–540 nm) filters. The DAF-2 fluorescence images were analyzed with Argus-100 imaging analysis system (Hamamatsu Photonics, Hamamatsu, Japan). Changes in the cellular DAF-2 fluorescence intensities (F) in each experiment were normalized to the baseline fluorescence recorded prior to stimulation (F0). Changes in NO production are expressed as F/F0, thus representing percentage increases above basal levels. In some experiments, L-NG-nitroarginine methyl ester (L-NAME) and DNAME were added 15 min before loading with DAF-2 DA. Because NOS generates O2− instead of NO in the absence of Larginine (Xia et al., 1998), we added 100 μM L-arginine to PSS for

Fig. 9 – Immunofluorescent staining of nNOS and P2X2 in an isolated OHC. (A) Immunofluorescent staining for nNOS in an isolated OHC. Note that intense staining of nNOS was seen in the apical region of IHC including hair bundles and the cuticular plate. No immunoreactivity was seen in the control OHCs either without nNOS primary antibody or using anti-nNOS blocking peptide. Scale bar, 10 μm. (B) Double immunofluorescent staining for nNOS and P2X2 in an isolated OHC. Immunofluorescent staining for nNOS was visualized with rhodamine (red), while P2X2 immunofluorescence was visualized with FITC (green). Note that co-localization of P2X2 and nNOS was confined to the apical region of OHCs. Scale bar, 10 μm. (C). Western blot analysis confirms the expression of nNOS and P2X2 in cochleae of guinea pigs. 155-kDa and 64-kDa bands are revealed in organ of Corti homogenate probed with nNOS and P2X2 antiserum, respectively.

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accomplished using a Fluoview FV300 confocal laser scanning microscope with a 488-nm line of an argon ion laser using 20× water immersion lens (Olympus, Tokyo, Japan). Images of single OHC were magnified 5× digitally. Background fluorescence was measured and subtracted from the signal to obtain the fluorescence intensity value. Changes in the DAF-2 fluorescence over time were analyzed offline using the Fluoview software (Olympus, Tokyo, Japan). The images were then digitally inverted to increase the contrast and for clarity. All images were ultimately prepared using Adobe Photoshop™ software.

4.4. Simultaneous measurement of [Ca2+]i and NO production Isolated OHCs were incubated simultaneously with 2 μM fura2/AM and 5 μM DAF-2 DA for 30 min at room temperature. Fluorescence was excited by alternately illuminating the OHCs at 340 nm and 360 nm for the fura-2 measurements and 485– 495 nm for the DAF-2 measurements.

4.5.

Measurements of Mn2+ influx

The Mn2+ quenching fluorescence signal was monitored using the same equipment as that for the [Ca2+]i measurements. An excitation wavelength of 360 nm was used since the intracellular changes in Ca2+ were insensitive at this wavelength. To confirm the participation of the solely Mn2+ influx, CaCl2 was omitted in PSS. 50 μM Mn2+ was added to the cell perfusion solution 4 min after ATP stimulation. L-NAME and SNAP were added before starting the experiments. To obtain the background fluorescence intensity, 0.1% Triton X-100 was added at the end of each experiment. Thereafter, maximal Mn2+ quenching value was estimated by subtraction of the background fluorescence intensity. The data were expressed as the % decrease of F360.

4.6. Fig. 10 – Spatiotemporal distribution of NO production induced by ATP. (A) Local application of ATP to different regions of an isolated OHC via a puff pipette. The locations of the tip of the pipette are illustrated in the left panel. Substantial ATP-induced NO production was observed with the pipette placed at the apex of the cell (upper trace). Local application of PSS to each region of the OHC did not induce NO production (not shown). (B) Pseudocolor images of NO production during ATP stimulation in real-time recording using confocal laser microscopy. Application of 100 μM ATP to the perfusion solution induced an increase in DAF-2 fluorescence, which initially occurred in the apical region. Note that no detectable NO production was observed in the basal region in this cell during ATP stimulation.

DAF-2 measurements except for the experiments using L-NAME and D-NAME. Control perfusion of PSS with 100 μM L-arginine did not induce any changes in DAF-2 fluorescence in the present study, which corresponds with the findings of a previous study (Kimura et al., 2004). For the temporal and spatial analysis of the ATP-induced NO production, confocal imaging of DAF-2 loaded OHCs was

Immunohistochemistry

The cellular localization of the nNOS isoforms in isolated OHCs was analyzed by immunofluorescent staining. Isolated OHCs were transferred into a Lab-Tek chamber slide (Lab-Tek, Nalge Nunc International, NY, USA) and fixed with buffered 4% paraformaldehyde for 30 min at room temperature. Then the cells were immersed in 0.3% triton X-100 in PBS for 5 min and incubated with blocking solution (10% normal goat serum and 1% bovine serum albumin with 0.3% triton X-100 in PBS) for 30 min at room temperature. Thereafter, a polyclonal antibody against a peptide corresponding to residues 1095–1289 of rabbit nNOS (BD Transduction Laboratories, CA, USA) was used at 1:250 dilution for 2–2.5 h at 37 °C. nNOS antibody was visualized by incubation for 45 min at room temperature with FITCconjugated anti-rabbit IgG (Jackson ImmunoResearch Laboratories) diluted 1:100 in PBS. Negative controls were processed with an omission of either primary antibodies or primary antibody adsorbed with an excess concentration of anti-nNOS blocking peptide (BD Biosciences, MA, USA). No detectable labeling was observed in either control. For the double staining of nNOS and P2X2 in isolated OHCs, polyclonal rabbit anti P2X2 (a peptide corresponding to residues 457–472 of P2x2, 1:200, Alomone Labs, Jerusalem, Isrreal) and

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polyclonal mouse anti-nNOS (a peptide corresponding to residues 1095–1289 of nNOS, 1:250; BD transduction Laboratories, CA) were used as primary antibodies. FITC-conjugated donkey anti-rabbit IgG (1:100) and CYTM3-conjugated goat antimouse IgG (1:200, Jackson ImmunoRsearch laboratory) were used to detect the labeling of P2X2 and nNOS, respectively. Negative controls were processed by omitting either the primary antibodies for nNOS and P2X2 or the secondary antibodies. Immunofluorescent staining of isolated OHCs was examined under confocal laser scanning microscopy (Olympus Fluoview, BX-50, Japan). This instrument was equipped with the appropriate emission filter for FITC (488 nm), for Cy3 (568 nm), and was controlled by a computer via an ultrafast SCSI interface. Fluorescent images 600 × 800 pixels were processed and stored using the FLUOVIEW FV300 software (Olympus, Japan).

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puff ejector (FemtoJet, Eppendorf, 10 s pulses, pipette tip diameter of 2–5 μm) and micromanipulator (model 5171, Eppendorf). The puffer pipettes were held about 10–20 μm away from OHCs. All experiments were performed at room temperature (20–24 °C).

4.9.

Statistical analysis

All the values are given as the means ± standard deviation of the mean, and n denotes the number of the cells per experiment. Statistical significance was evaluated by one-way ANOVA for comparison between different treatments and by paired t test for comparison between the first and secondary calcium response. P values less than 0.05 are considered to be significant.

Acknowledgment 4.7.

Western blot analysis

Western blot analysis was used to confirm the specificity of immunostaining for nNOS and P2X2 in OHCs of guinea pigs. Organ of Corti from 24 cochleae of guinea pigs was homogenized and centrifuged at 10,000 × g for 15 min at 4 °C. Protein was determined in the resulting supernatant fractions using a BioRad DC protein assay kit (Bio-Rad Laboratories, CA), and aliquots were stored at −80 °C until for use. After denatured the sample proteins at 100 °C for 5 min in SDS sample buffer (3:1), 20 μg of proteins was loaded and separated on 7.5% SDS-PAGE, then transferred to a PVDF membrane (Bio-Rad, Hercules CA). Immunoblots were performed with polyclonal rabbit antibody against nNOS (1:200, BD Transduction Laboratories, CA) and rabbit antibody against P2X2 (1:200, Alomone Labs, Jerusalem, Israel), respectively, followed by incubation with anti-rabbit horseradish peroxidase-conjugated secondary antibody (1:2000, Amersham Biosciences), and finally visualized by enhanced chemiluminescence (ECL detection reagents, Amersham Biosciences). Because rabbit and mouse anti nNOS are corresponding to the same residues of nNOS (1095–1289), only rabbit anti nNOS was detected in the present study.

4.8.

Drugs and solutions

All drugs were purchased from Sigma, unless otherwise specified. KT5823 was obtained from Calbiochem (La Jolla, CA, USA), and 1-(2-trifluoromethylphenyl) imidazole (TRIM) was from ALEXIS Biochemicals. Ca2+-free solution was prepared by omitting CaCl2 from PSS and by adding 2 mM of ethylene glycolbis (β-aminoethyl ether) N,N,N′,N′-tetraacetic acid. 7-Nitroindazole (7-NI), S-nitro-N-acetylpenicillamine (SNAP), TRIM, KT5823 and oxadiazolo(4,3-a)quinoxalin-1-one (ODQ) were dissolved in dimethylsulfoxide (DMSO) and stored as a stock solution. Prior to the experiment each day, final solutions were prepared by dissolving the stock solution in PSS. The final concentrations of DMSO did not exceed 0.01%. The concentrations of DMSO (b0.01%) used in the present study had no significant effect on the [Ca2+]i of the cells. During the experiment, the recording chamber was continuously perfused with PSS at 400 μl/min using a micro-perfusion system (microperpex, LKB2132, Pharmacia LKB Biotechnology, Uppsala, Sweden). Each substance dissolved in PSS was usually applied by perfusion. For the local application of ATP, ATP dissolved in PSS was applied by using a

We thank Dr. Masahiro Oike for the critical comments and helpful suggestions regarding the DAF-2 measurements.

REFERENCES

Ashmore, J.F., Ohmori, H., 1990. Control of intracellular calcium by ATP in isolated outer hair cells of the guinea-pig cochlea. J. Physiol. 428, 109–131. Ayajiki, K., Fujioka, H., Okamura, T., Toda, N., 2001. Relatively selective neuronal nitric oxide synthase inhibition by 7-nitroindazole in monkey isolated cerebral arteries. Eur. J. Pharmacol. 423, 179–183. Boulton, C.L., Southam, E., Garthwaite, J., 1995. Nitric oxide-dependent long-term potentiation is blocked by a specific inhibitor of soluble guanylyl cyclase. Neuroscience 69, 699–703. Breer, H., Shepherd, G.M., 1993. Implications of the NO/cGMP system for olfaction. Trends Neurosci. 1, 5–9. Broillet, M.C., Randin, O., Chatton, J.Y., 2001. Photoactivation and calcium sensitivity of the fluorescent NO indicator 4-5,-diaminofluorescein (DAF-2): implications for cellular NO imaging. FEBS Lett. 491, 227–232. Burkhardt, M., Glazova, M., Gambaryan, S., Vollkommer, T., Butt, E., Bader, B., Heermeier, K., Lincoln, T.M., Walter, U., Palmetshofer, A., 2000. KT5823 inhibits cGMP-dependent protein kinase activity in vitro but not in intact human platelets and rat mesangial cells. J. Biol. Chem. 275, 33536–33541. Cho, H., Harada, N., Yamashita, T., 1997. Extracellular ATP-induced Ca2+ mobilization of type I spiral ganglion cells from the guinea pig cochlea. Acta Otolaryngol. 117, 545–552. Collmann, C., Carlsson, M.A., Hansson, B.S., Nighorn, A., 2004. Odorant-evoked nitric oxide signals in the antennal lobe of Manduca Sexta. J. Neurosci. 24, 6070–6077. Cudeiro, J., Rivadulla, C., 1999. Sight and insight—On the physiological role of nitric oxide in the visual system. Trends Neurosci. 22, 109–116. Dallos, P., 1992. The active cochlea. J. Neurosci. 12, 4575–4585. Dallos, P., Corey, M.E., 1991. The role of outer hair cell motility in cochlear tuning. Curr. Opin. Neurobiol. 1, 215–220. Dedkova, E.N., Blatter, L.A., 2002. Nitric oxide inhibits capacitative Ca2+ entry and enhances endoplasmic reticulum Ca2+ uptake in bovine vascular endothelial cells. J. Physiol. 539, 77–91. Dumont, R.A., Lins, U., Filoteo, A.G., Penniston, J.T., Kachar, B., Gillespie, P.G., 2001. Plasma membrane Ca2+-ATPase isoform 2a is the PMCA of hair bundles. J. Neurosci. 21, 5066–5078. Esplugues, J.V., 2002. NO as a signalling molecule in the nervous system. Br. J. Pharmacol. 135, 1079–1095.

112

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Franz, P., Hauser-Kronberger, C., Bock, P., Quint, C., Baumgartner, W.D., 1996. Localization of nitric oxide synthase I and III in the cochlea. Acta Otolaryngol. 116, 726–731. Frolenkov, G.I., Mammano, F., Kachar, B., 2003. Regulation of outer hair cell cytoskeletal stiffness by intracellular Ca2+: underlying mechanism and implications for cochlear mechanics. Cell Calcium 33, 185–195. Furness, D.N., Karkanevatos, A., West, B., Hackney, C.M., 2002. An immunogold investigation of the distribution of calmodulin in the apex of cochlear hair cells. Hear. Res. 173, 10–20. Garcia-Pascual, A., Costa, G., Labadia, A., Jimenez, E., Triguero, D., 1999. Differential mechanisms of urethral smooth muscle relaxation by several NO donors and nitric oxide. Naunyn-Schmiedeberg's Arch. Pharmacol. 360, 80–91. Gosepath, K., Gath, I., Maurer, J., Pollock, J.S., Amedee, R., Forstermann, U., Mann, W., 1997. Characterization of nitric oxide synthase isoforms expressed in different structures of the guinea pig cochlea. Brain Res. 747, 26–33. Grynkiewicz, G., Poenie, M., Tsien, R.Y., 1985. A new generation of Ca2+ indicators with greatly improved fluorescence properties. J. Biol. Chem. 260, 3440–3450. Handy, R., Moore, P., 1997. Mechanism of the inhibition of neuronal nitric oxide synthase by 1-(2-trifluoromethylphenyl) imidazole (TRIM). Life Sci. 60, 389–394. Harada, N., Ernst, A., Zenner, H.P., 1993. Hyposmotic activation hyperpolarizes outer hair cell of guinea pig cochlea. Brain Res. 613, 205–211. He, D.Z., Jia, S., Dallos, P., 2004. Mechanoelectrical transduction of adult outer hair cells studied in a gerbil hemicochlea. Nature 429, 766–770. Heinrich, U., Maurer, J., Koesling, D., Mann, W., Forstermann, U., 2000. Immuno-electron microscopic localization of the alpha(1) and beta(1)-subunits of soluble guanylyl cyclase in the guinea pig organ of Corti. Brain Res. 885, 6–13. Heinrich, U.R., Maurer, J., Mann, W., 2004. Evidence for a possible NOS back-up system in the organ of Corti of the guinea pig. Eur. Arch. Oto-Rhino-Laryngol. 261, 121–128. Housley, G.D., Kanjhan, R., Raybould, N.P., Greenwood, D., Salih, S. G., Jarlebark, L., Burton, L.D., Setz, V.C., Cannell, M.B., Soeller, C., Christie, D.L., Usami, S., Matsubara, A., Yoshie, H., Ryan, A. F., Thorne, P.R., 1999. Expression of the P2X(2) receptor subunit of the ATP-gated ion channel in the cochlea: implications for sound transduction and auditory neurotransmission. J. Neurosci. 19, 8377–8388. Housley, G.D., Jagger, D.J., Greenwood, D., Raybould, N.P., Salih, S. G., Jarlebark, L.E., Vlajkovic, S.M., Kanjhan, R., Nikolic, P., Munoz, D.J., Thorne, P.R., 2002. Purinergic regulation of sound transduction and auditory neurotransmission. Audiol. Neurootol. 7, 55–61. Kennedy, H.J., Crawford, A.C., Fettiplace, R., 2005. Force generation by mammalian hair bundles support a role in cochlear amplification. Nature 433, 880–883. Kimura, C., Oike, M., Ohnaka, K., Nose, Y., Ito, Y., 2004. Constitutive nitric oxide production in bovine aortic and brain microvascular endothelial cells: a comparative study. J. Physiol. 554, 721–730. Kojima, H., Nakatsubo, N., Kikuchi, K., Kawahara, S., Kirino, Y., Nagoshi, H., Hirata, Y., Nagano, T., 1998. Detection and imaging of nitric oxide with novel fluorescent indicators: diaminofluoresceins. Anal. Chem. 70, 2446–2453. Li, N., Sul, J.Y., Haydon, P.G., 2003. A calcium-induced calcium influx factor, nitric oxide, modulates the refilling of calcium stores in astrocytes. J. Neurosci. 23, 10302–10310. Lin, S., Fagan, K.A., Li, K.X., Shaul, P.W., Cooper, D.M., Rodman, D. M., 2000. Sustained endothelial nitric-oxide synthase activation requires capacitative Ca2+ entry. J. Biol. Chem. 275, 17979–17985. Mammano, F., Frolenkov, G.I., Lagostena, L., Belyantseva, I.A., Kurc, M., Dodane, V., Colavita, A., Kachar, B., 1999. ATP-induced Ca2+ release in cochlear outer hair cells: localization of an

inositol triphosphate-gated Ca2+ store to the base of the sensory hair bundle. J. Neurosci. 19, 6918–6929. Raybould, N.P., Housley, G.D., 1997. Variation in expression of the outer hair cell P2X receptor conductance along the guinea-pig cochlea. J. Physiol. 498, 717–727. Robles, L., Ruggero, M.A., 2001. Mechanics of the mammalian cochlea. Physiol. Rev. 81, 1305–1352. Sagami, I., Daff, S., Shimizu, T., 2001. Intra-subunit and inter-subunit electron transfer in neuronal nitric-oxide synthase: effect of calmodulin on heterodimer catalysis. J. Biol. Chem. 276, 30036–30042. Scrogin, K.E., Hatton, D.C., Chi, Y., Luft, F.C., 1998. Chronic nitric oxide inhibition with L-NAME: effects on autonomic control of cardiovascular system. Am. J. Physiol. 274, R367–R374. Shen, J., Harada, N., Yamashita, T., 2003. Nitric oxide inhibits adenosine 5′-triphosphate-induced Ca2+ response in inner hair cells of the guinea pig cochlea. Neurosci. Lett. 337, 135–138. Shen, J., Harada, N., Nakazawa, H., Yamashita, T., 2005. Involvement of the nitric oxide/cyclic GMP pathway and neuronal nitric oxide synthase in ATP-induced Ca2+ signalling in cochlear inner hair cells. Eur. J. Neurosci. 21, 2912–2922. Shi, X., Ren, T., Nuttall, A.L., 2001. Nitric oxide distribution and production in the guinea pig cochlea. Hear. Res. 153, 23–31. Sugasawa, M., Erostegui, C., Blanchet, C., Dulon, D., 1996. ATP activates non-selective cation channels and calcium release in inner hair cells of the guinea-pig cochlea. J. Physiol. 491, 707–718. Suzuki, N., Kojima, H., Urano, Y., Kikuchi, K., Hirata, Y., Nagano, T., 2002. Orthogonality of calcium concentration and ability of 4,5-diaminofluorescein to detect NO. J. Biol. Chem. 277, 47–49. Szonyi, M., He, D.Z., Ribari, O., Sziklai, I., Dallos, P., 2001. Intracellular calcium and outer hair cell electromotility. Brain Res. 922, 65–70. Takumida, M., Anniko, M., 2001. Detection of nitric oxide in the guinea pig inner ear, using a combination of aldehyde fixative and 4,5-diaminofluorescein diacetate. Acta Oto-Laryngol. 121, 460–464. Takumida, M., Anniko, M., 2002. Nitric oxide in the inner ear. Curr. Opin. Neurol. 15, 11–15. Takumida, M., Anniko, M., Popa, R., Zhang, D.M., 2000. Localization of soluble guanylate cyclase activity in the guinea pig inner ear. Acta Oto-Laryngol. 120, 28–33. Tsumamoto, Y., Yamashita, K., Takumida, M., Okada, K., Mukai, S., Shinya, M., Yamashita, H., 2002. In situ localization of nitric oxide synthase and direct evidence of NO production in rat retinal ganglion cells. Brain Res. 933, 118–129. Van den Abbeele, T., Tran Ba Huy, P., Teulon, J., 1996. Modulation by purines of calcium-activated non-selective cation channels in the outer hair cells of the guinea-pig cochlea. J. Physiol. 494, 77–89. Wang, G.Y., Liets, L.C., Chalupa, L.M., 2003. Nitric oxide differentially modulates ON and OFF responses of retinal ganglion cells. J. Neurophysiol. 90, 1304–1313. Xia, Y., Tsai, A.L., Berka, V., Zweier, J.L., 1998. Superoxide generation from endothelial nitric-oxide synthase. A Ca2+/ calmodulin-dependent and tetrahydrobiopterin regulatory process. J. Biol. Chem. 273, 25804–25808. Yamashita, T., Amano, H., Kumazawa, T., 1993. Efferent neurotransmitters and intracellular Ca2+ concentrations in inner hair cells of guinea pig. ORL J. Otorhinolaryngol. Relat. 55, 201–204. Yao, S.T., Gourine, A.V., Spyer, K.M., Barden, J.A., Lawrence, A.J., 2003. Localization of P2X2 receptor subunit immunoreactivity on nitric oxide synthase expressing neurons in the brain stem and hypothalamus of the rat: a fluorescence immunohistochemical study. Neuroscience 121, 411–419. Yukawa, H., Shen, J., Harada, N., Cho-Tamaoka, H., Yamashita, T., 2005. Acute effects of glucocorticoids on ATP-induced Ca2+ mobilization and nitric oxide production in cochlear spiral ganglion neurons. Neuroscience 130, 485–496.