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The Journal of Physiology logoLink to The Journal of Physiology
. 2004 Aug 26;560(Pt 3):737–752. doi: 10.1113/jphysiol.2004.069294

Neuroepithelial oxygen chemoreceptors of the zebrafish gill

Michael G Jonz 1, Ian M Fearon 1, Colin A Nurse 1
PMCID: PMC1665292  PMID: 15331683

Abstract

In aquatic vertebrates, hypoxia induces physiological changes that arise principally from O2 chemoreceptors of the gill. Neuroepithelial cells (NECs) of the zebrafish gill are morphologically similar to mammalian O2 chemoreceptors (e.g. carotid body), suggesting that they may play a role in initiating the hypoxia response in fish. We describe morphological changes of zebrafish gill NECs following in vivo exposure to chronic hypoxia, and characterize the cellular mechanisms of O2 sensing in isolated NECs using patch-clamp electrophysiology. Confocal immunofluorescence studies indicated that chronic hypoxia (PO2 = 35 mmHg, 60 days) induced hypertrophy, proliferation and process extension in NECs immunoreactive for serotonin or synaptic vesicle protein (SV2). Under voltage clamp, NECs responded to hypoxia (PO2 = 25–140 mmHg) with a dose-dependent decrease in K+ current. The current–voltage relationship of the O2-sensitive current (IKO2) reversed near EK and displayed open rectification. Pharmacological characterization indicated that IKO2 was resistant to 20 mm tetraethylammonium (TEA) and 5 mm 4-aminopyridine (4-AP), but was sensitive to 1 mm quinidine. In current-clamp recordings, hypoxia produced membrane depolarization associated with a conductance decrease; this depolarization was blocked by quinidine, but was insensitive to TEA and 4-AP. These biophysical and pharmacological characteristics suggest that hypoxia sensing in zebrafish gill NECs is mediated by inhibition of a background K+ conductance, which generates a receptor potential necessary for neurosecretion and activation of sensory pathways in the gill. This appears to be a fundamental mechanism of O2 sensing that arose early in vertebrate evolution, and was adopted later in mammalian O2 chemoreceptors.

Regulation of arterial oxygen levels within a normal physiological range during periods of low O2 exposure, or hypoxia, is of central importance to the survival of cells and tissues. In mammals, specialized cells sensitive to changes in O2 tension (PO2) initiate local or centrally mediated physiological adjustments. These cells include type I cells of the carotid body (González et al. 1994; Peers, 1997; López-Barneo et al. 2001), neuroepithelial bodies (NEBs) of the lung (Youngson et al. 1993; Fu et al. 2002), adrenal chromaffin cells (Thompson & Nurse, 1998), central (Plant et al. 2002; Neubauer & Sunderram, 2004) and peripheral (Campanucci et al. 2003) neurones, and vascular smooth muscle cells (Michelakis et al. 1995). The initial transduction step in most O2-sensitive cells involves modulation of K+ conductance and, in peripheral chemoreceptors such as type I cells and NEBs, may lead to Ca2+-dependent neurosecretion and activation of postsynaptic pathways (González et al. 1994; López-Barneo et al. 2001; Fu et al. 2002).

Recently, a group of background or ‘leak’ K+ channels have been identified as important targets for hypoxic modulation in a variety of O2-sensitive cells, where they determine or contribute to the receptor potential (Buckler, 1997; Buckler et al. 2000; Hartness et al. 2001; Plant et al. 2002; Campanucci et al. 2003; Kemp et al. 2004). These two-pore (2P) domain K+ channels are voltage-independent and play an important role in setting the resting membrane potential and input resistance of the cell (Goldstein et al. 2001; Lesage, 2003). In most cases, the O2-sensitive background K+ channels are inhibited by hypoxia and are resistant to conventional blockers of voltage-dependent K+ channels, such as tetraethylammonium (TEA) and 4-aminopyridine (4-AP), but are quite sensitive to quinidine (Buckler, 1997; O'Kelly et al. 1999; Buckler et al. 2000; Campanucci et al. 2003).

In addition to the well-characterized electrophysiological responses to acute hypoxia, significant morphological changes in O2 chemoreceptors have been attributed to prolonged, or chronic, exposure to hypoxia. Cellular hypertrophy and proliferation occurred in type I chemoreceptor cells after in vitro (Stea et al. 1992; Mills & Nurse, 1993; Nurse & Vollmer, 1997) and in vivo (for review see Wang & Bisgard, 2002) exposure to hypoxia, and this may contribute to enlargement of the carotid body during acclimatization to hypoxia.

Characterization of physiological and morphological responses of peripheral O2 chemoreceptors to hypoxia at the cellular level has, so far, been confined to those of mammals. However, work on aquatic vertebrates, such as teleost fish, may offer certain advantages. Fish naturally experience fluctuations in water PO2 and are known to exhibit pronounced responses to hypoxia, such as hyperventilation, bradycardia and changes in gill vascular resistance (for review see Burleson et al. 1992). These responses appear to arise principally from peripheral O2 chemoreceptors of the gills (Milsom & Brill, 1986; Burleson et al. 1992).

In vertebrates, the pharyngeal arches give rise to important respiratory structures, such as the carotid body in mammals and the gills in fish (Weichert, 1967). In teleost fish, four arches are innervated by branches of the glossopharyngeal (first arch only) and vagus nerves, and bear numerous gill filaments with respiratory lamellae (Fig. 1A). Secretory neuroepithelial cells (NECs) are similar in morphology to mammalian O2 chemoreceptors and have been identified in the gill filaments of all fish species examined (for references see Jonz & Nurse, 2003). NECs reside within the filament epithelium, between the incident flow of water through the gills and the arterial blood supply (Fig. 1B). Thus, NECs are in a favourable position to detect changes in water or arterial PO2. In a recent study, we characterized the morphology, organization and innervation patterns of NECs in whole-mount preparations of the adult zebrafish gill (Jonz & Nurse, 2003). NECs were present on all four gill arches, and two neural pathways were identified that may be involved in hypoxic signalling and initiation of the hypoxia response in fish (Fig. 1B; Jonz & Nurse, 2003). Furthermore, many NECs contain the neurotransmitter serotonin (5-HT) and synaptic vesicles are polarized within the basal cytoplasm near adjacent nerve fibres (Dunel-Erb et al. 1982; Jonz & Nurse, 2003). At the ultrastructural level, these vesicles appeared to degranulate in response to acute hypoxia (Dunel-Erb et al. 1982), suggesting that NECs may have a chemosensory role in the gill.

Figure 1. Simplified illustration of the position of neuroepithelial cells (NECs) within the gill filaments of zebrafish.

Figure 1

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A, distal section of a single gill filament (F) with perpendicular respiratory lamellae (L). Large arrows indicate the incident flow of water that is diverted by the filament epithelium and flows over the lamellae during ventilation. Small arrows indicate the flow of blood distally through the afferent filament artery (aFA) before oxygenation, and the flow of oxygen-rich blood through the lamellae and efferent filament artery (eFA). B, summary diagram showing the major structures involved in potential O2-sensing pathways of the zebrafish gill. Transverse section of the filament depicted in A showing a NEC residing within the filament epithelium (FE) in close proximity to the respiratory water flow (large arrows) and the eFA. Dashed arrows indicate possible routes through which hypoxia may be detected by NECs. NECs receive innervation from a nerve plexus (NP) arising from the extrinsic nerve bundle (eNB) and from fibres of the intrinsic nerve bundle (iNB). In addition, NECs make contact with intrinsic fibres via neurone-like processes. Based on observations by Jonz & Nurse (2003).

In this study, we used the zebrafish gill preparation to provide the first functional characterization of a peripheral O2 chemoreceptor at the cellular level in an aquatic vertebrate. The zebrafish may be an important animal model for further studies on the effects of developmental and genetic manipulations on O2 chemoreception in vertebrates. We observed pronounced morphological changes in gill NECs in whole-mount preparations following in vivo exposure to chronic hypoxia. In addition, a K+ conductance that was sensitive to acute hypoxia, leading to membrane depolarization, was identified in isolated NECs. This work reveals the O2-sensitivity of gill NECs and suggests that they may indeed function as peripheral O2 chemoreceptors in fish.

Methods

Animals

Adult zebrafish (150–400 mg) were obtained from local commercial sources and maintained at 28°C on a 14 h light–dark cycle (Westerfield, 2000). All procedures for animal use were carried out according to institutional guidelines, which adhere to those of the Canadian Council on Animal Care (CCAC).

Chronic hypoxia

The effects of prolonged exposure to hypoxia on neuroepithelial cell (NEC) morphology were examined. Zebrafish were exposed to chronic hypoxia in vivo at 28°C in a 4-l aquarium in which water PO2 was controlled using a mixture of compressed air and 100% N2. Animals were slowly acclimated to hypoxia by gradually decreasing water PO2 from 150 to 35 mmHg in intervals of ∼30 mmHg every 3 or 4 days over a 14-day period. Zebrafish were subsequently exposed to a PO2 of 35 mmHg for a further 46 days (total exposure, 60 days). This O2 tension was above the critical PO2 for adult zebrafish (20 mmHg at 28°C; Barrionuevo & Burggren, 1999) and was selected to minimize mortality while maximizing the morphological changes in NECs associated with chronic hypoxia. Control zebrafish were maintained under similar conditions, but with continuously aerated water.

Confocal immunofluorescence

Techniques for tissue extraction, immunolabelling and confocal imaging were performed as previously described (Jonz & Nurse, 2003). Zebrafish were killed by overdose with 1 mg ml−1 MS 222 (tricaine, Sigma) and decapitated. Whole gill baskets were removed and rinsed in ice-cold phosphate-buffered solution (PBS) containing (mm): NaCl 137, Na2HPO4 15.2, KCl 2.7 and KH2PO4 1.5; pH 7.8 (Bradford et al. 1994). Gill baskets were fixed by immersion in 4% paraformaldehyde in PBS at 4°C overnight. Tissue was rinsed in PBS and permeabilized for 48–72 h at 4°C. Permeabilizing solution (PBS-TX) contained 1% fetal calf serum (FCS) and 0.5% Triton X-100 in PBS (pH 7.8). Gill arches were then separated and prepared individually as whole mounts for immunocytochemistry. Neuroepithelial cells (NECs) of the gill filaments were identified in whole-mount preparations using antisera against serotonin (5-HT) and the synaptic vesicle protein SV2 (Dunel-Erb et al. 1982; Jonz & Nurse, 2003). NECs of zebrafish display SV2-immunoreactivity (IR), most of which are also serotonergic (Jonz & Nurse, 2003). Extrinsic nerve fibres were identified using antiserum against a zebrafish-derived neurone-specific antigen (zn-12). zn-12 antiserum acts as a general neuronal marker in zebrafish (Trevarrow et al. 1990), and its labelling of neural structures of the gill has been previously characterized (Jonz & Nurse, 2003). Polyclonal rabbit anti-5-HT (Sigma) was used at a dilution of 1 : 200 and localized with goat anti-rabbit secondary antiserum conjugated with the green flourophore, flourescein isothiocyanate (FITC, 1 : 50, Jackson ImmunoResearch Laboratories Inc., West Grove, PA, USA). Monoclonal mouse anti-zn-12 and anti-SV2 (Developmental Studies Hybridoma Bank, University of Iowa, IA, USA) were used at a dilution of 1 : 100 and 1 : 200, respectively, and localized with goat anti-mouse secondary antiserum conjugated with the red fluorophore, Alexa 594 (1 : 100, Molecular Probes, Eugene, OR, USA). All antisera were diluted with PBS-TX. Whole gill arches were incubated in primary antisera for 24–48 h at 4°C. Tissue was rinsed in PBS-TX and treated with secondary antisera at room temperature (22–24°C) for 1 h in darkness. Arches were rinsed in PBS and mounted on glass microscope slides in Vectashield (Vector Laboratories Inc., Burlingame, CA, USA) to reduce photobleaching. Whole-mount preparations were examined using an upright microscope (Eclipse E800, Nikon) and a confocal scanning system (Radiance 2000, Bio-Rad, Hercules, CA, USA) equipped with argon (Ar) and helium–neon (He–Ne) lasers with peak outputs of 488 nm and 543 nm, respectively. Images were detected with a photomultiplier tube and photodiode array, and were collected using confocal graphics software (LaserSharp 2000, Bio-Rad). Each image is presented as a composite projection of serial optical sections. Image processing and manipulation was performed using Corel Draw 9 (Corel Corp., Ottawa, ON, Canada).

Morphometric analysis

Confocal images of NECs from whole-mount gill preparations of zebrafish exposed to normoxia (Nox, control) and chronic hypoxia (CHox) were examined using analytical imaging software (Laser Pix, Bio-Rad). All NECs contained within a 150 µm × 75 µm sampling area from the distal region of a single gill filament in each animal were included in the analysis. Each sample represents the average value from NECs located within the sampling area of a single animal. Manual threshold adjustment for each image permitted automated collection of data. Two-dimensional or projection area (µm2), proportional to cell size, was measured in NECs of Nox and CHox fish. In addition, attributes such as number, aspect ratio (ratio of length to width) and shape factor of 5-HT-IR NECs were compared in both groups. Shape factor (SF) was calculated according to the equation SF = 4π A P−2, where A is area and P is perimeter. Shape factor analysis allows measurement of changes in complexity of shape and membrane surface during cellular growth and differentiation (Levallois et al. 1997), where values approaching zero indicate a greater complexity of shape (i.e. less round). Changes in the number of SV2-IR NECs were also observed. These cells included both 5-HT-positive and 5-HT-negative NECs. In addition, the number of 5-HT-IR NECs having neurone-like processes extending towards intrinsic nerve fibres, and process length, were compared in zebrafish exposed to Nox and CHox.

Cell isolation

Zebrafish were stunned with a blow to the head and quickly decapitated. All procedures for the dissociation and isolation of NECs were carried out under sterile conditions in a laminar flow hood. Gill baskets were removed and rinsed in a wash solution (2% penicillin/streptomycin in PBS) for 10 min. All four gill arches were then separated and distal filament regions, areas rich in NECs (Jonz & Nurse, 2003), were selectively removed. Tissue was placed in 0.25% trypsin/EDTA for 1 h at room temperature, minced with fine forceps and triturated in a centrifuge tube with a Pasteur pipette. The trypsin reaction was stopped with the addition of 10% FCS. Lower concentrations of trypsin or mechanical separation alone did not permit adequate dissociation. The cell suspension was centrifuged (140 g) for 3 min and the pellet triturated in PBS. Gill cells were centrifuged once more and suspended in Leibovitz's (L-15, with l-glutamine, Gibco, Grand Island, NY, USA) culture medium supplemented with 1% penicillin/streptomycin and 5% FCS. Cells were plated in 0.1 ml L-15 to promote attachment to the central well of modified culture dishes, and 2 ml L-15 was added after 18–24 h. Polystyrene culture dishes (Falcon) were modified by drilling a small central hole in the bottom and attaching a glass coverslip to the underside with Sylgard (Dow Corning Corp., Midland, MI, USA). Dishes with glass-bottom wells were UV-sterilized and coated with poly l-lysine (0.1 mg ml−1) and Matri-Gel (Becton Dickenson, Mississauga, ON, Canada) following manufacturer specifications. NECs that adhered to the culture substrate were identified using 2 mg ml−1 Neutral Red (NR, Sigma), a vital marker used to identify 5-HT-containing cells (Stuart et al. 1974; Youngson et al. 1993). To verify the staining of NECs by NR, vital staining of dissociated gill cells was followed by fixation with 4% paraformaldehyde in PBS for 15 min at room temperature, and immunolabelling with 5-HT antiserum, as described above. Cells were observed with an inverted microscope (IM-35, Zeiss) equipped with epi-fluorescence, and images were captured with a digital camera (Retiga, QImaging, Burnaby, BC, Canada) and imaging software (Northern Eclipse, Empix Imaging Inc., Mississauga, ON, Canada). For these experiments, cells were plated in dishes fitted with coverslips etched with grids (Eppendorf, Hamburg, Germany) to facilitate location of NR-positive 5-HT-IR cells (NECs) from digital images. Serotonergic NECs of the gill in zebrafish are found in both the filament epithelium and respiratory lamellae (Jonz & Nurse, 2003). To further aid in differentiating between NECs of the filament and the smaller NECs of the lamellae (∼5 µm), the diameter of dissociated 5-HT-IR NECs was measured in vitro. Based on these observations, NR-positive cells ≥ 7 µm in diameter were selected for electrophysiological recording. This procedure favoured recordings of larger NECs located in the filament epithelium.

Electrophysiology

Dishes with attached NECs 24–48 h after dissociation were transferred to a perfusion chamber mounted on the stage of an inverted microscope (Axiovert S 100, Zeiss). Whole-cell patch-clamp recordings (Hamill et al. 1981) were performed using patch electrodes fabricated from borosilicate glass (World Precision Instruments, Inc., Sarasota, FL, USA). Glass electrodes were pulled on a vertical pipette puller (PP-83, Narishige, Japan) with a tip resistance of 5–8 MΩ and fire-polished. Electrodes were filled with intracellular recording solution containing (mm): KCl 135, NaCl 5, CaCl2 0.1, EGTA 11, Hepes 10 and Mg-ATP 2; pH adjusted to 7.2 with KOH. With this solution, intracellular Ca2+ was clamped below 1 nm (pCa 9.1). Extracellular recording solution contained (mm): NaCl 135, KCl 5, CaCl2 2, MgCl2 2, glucose 10 and Hepes 10; pH adjusted to 7.4 with NaOH. Junction potentials of ∼5 mV were subtracted. Seal resistance was typically 5–20 GΩ and holding current was < 3 pA at −60 mV.

Cells were held at −60 mV and currents were evoked by changing the membrane potential from −100 mV to +50 mV over a period of 1 s following a ramp protocol. Currents were also evoked by step depolarizations to various test potentials (see figure legends) for 50 ms at a frequency of 0.1 Hz. In such cases, steady-state current was measured as the average current evoked between 40 and 49 ms of the voltage step. Membrane capacitance (Cm) was measured automatically by Pulse software (Heka Electronik, Lambrecht, Germany). In some experiments, the slope of the current–voltage (I–V) relationship between −50 mV and −70 mV was used to calculate input resistance (Rin) according to Ohm's Law: Rin = ΔV ΔI−1. Values for passive membrane properties are expressed as means ± s.e.m. Voltage-clamp protocols were performed using an EPC 9 amplifier (Heka Electronik) and digitally converted using an A/D converter with an ITC-16 interface and Pulse software (Heka Electronik). Current-clamp recordings (I = 0 pA) were performed using the same apparatus and solutions as described for voltage-clamp experiments. In some experiments, voltage changes evoked by injection of 10 pA of hyperpolarizing current for 1 s was measured and Rin was calculated, as above. Recordings were filtered at 5 kHz and digitized at 10 kHz. All data were analysed using Pulsefit software (Heka Electronik) and figures were arranged using Origin 7.0 (OriginLab Corp., Northampton, MA, USA). In some traces, high frequency noise was filtered using Origin software.

The recording chamber was continuously perfused under gravity (4 ml min−1) with extracellular solution at room temperature (22–24°C). Hypoxia (PO2 = 25 mmHg) was produced by bubbling solution in the perfusion reservoir with 100% N2 for 30 min prior to recording. Tubing used to transfer the perfusate to the recording chamber was gas impermeable (Tygon, Saint-Gobain Performance Plastics Corporation, Akron, OH, USA). The degree and time course of changes in PO2 in the recording chamber were measured under identical experimental conditions using a carbon fibre electrode (10 µm, Dagan Corporation, Minneapolis, MN, USA) and EPC 9 amplifier. Control experiments were performed in which the perfusate was bubbled with compressed air instead of N2.

In pharmacological experiments designed to characterize the O2-sensitive K+ current (IKO2), conventional blockers of K+ channels were used. All drugs were purchased from Sigma and were dissolved in the extracellular solution (pH adjusted to 7.4). TEA (20 mm) and 4-AP (5 mm) were used in combination to block voltage-gated K+ channels, and quinidine (1 mm) was used to confirm the presence of background K+ channels.

Statistics

Data are presented as mean ± s.e.m. values for NEC number, area, process number and process length in Nox and CHox groups and were compared using the Student's unpaired t test. Differences in ratios, such as aspect ratio and shape factor, were appropriately compared using the non-parametric Mann Whitney U test. Multiple comparisons were performed using ANOVA followed by the Bonferroni test. For voltage and current-clamp analyses, mean ± s.e.m. values for current density (pA pF−1), membrane potential (Vm) and Rin in normoxic and hypoxic groups were compared using the paired t test. In Fig. 5C, where currents were normalized (%), the Mann Whitney U test was used to compare outward current under normoxic and hypoxic conditions. Statistical analyses were performed using software packages Excel (Microsoft, Redmond, WA, USA) and Statistix 2.0 (Analytical Software, Tallahassee, FL, USA).

Figure 5. Whole-cell, voltage-clamp recording of an O2-sensitive current in isolated neuroepithelial cells (NECs) of the zebrafish gill.

Figure 5

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A, whole-cell recording from an O2-sensitive NEC showing the current–voltage (I–V) relationship during exposure to normoxia (Nox, 150 mmHg), hypoxia (Hox, 25 mmHg) and after recovery in normoxia (Rec). Currents were evoked by changing the voltage from − 100 mV to +50 mV, following a ramp protocol, from a holding potential of −60 mV. IK was variable in these cells (216 ± 74.8 pA at +30 mV, n = 10). Inset, average response of 10 cells to ramp depolarization over a limited range of potentials during normoxia (Nox) and after acute hypoxia (Hox). B, average I–V relationship of the O2-sensitive difference current (i.e. Nox – Hox) from 10 cells. The O2-sensitive current (continuous trace) reverses near EK (calculated EK = −83 mV) and is therefore carried predominantly by K+ ions (i.e. IKO2). IKO2 was variable in NECs (58.8 ± 20.2 pA at +30 mV, n = 10). Dashed curve indicates the I–V relationship as predicted by the Goldman–Hodgkin–Katz current equation. C, time–series plot of IK inhibition during bath-perfusion with hypoxic solution. Whole-cell currents were evoked every 10 s using voltage steps from −60 mV to +30 mV. Mean ± s.e.m. (n = 8) steady-state current was normalized (left axis) to the average current evoked during the first 60 s (control) of recording and is displayed as a function of time. Changes in PO2 (right axis) in the recording chamber were measured with a carbon fibre electrode (continuous trace) during perfusion with hypoxic solution under identical experimental conditions. IK inhibition began immediately after hypoxic solution was introduced and reached a plateau as the PO2 levelled off near 25 mmHg. Outward current at 25 mmHg was significantly lower than in normoxia (Mann Whitney U test, P < 0.01). Upper traces, examples of whole-cell currents from a single cell before, during and after application of hypoxia. D, relationship between PO2 and IK inhibition by hypoxia in NECs. Data points taken from C during changes in PO2 (25–150 mmHg) are shown with a line of best fit (continuous line). Mean ± s.e.m. IK inhibition was 16.6 ± 2.5% at 25 mmHg. Dashed line shows the hypoxia-induced increase in nerve discharge (normalized; right axis) recorded from sensory fibres of the glossopharyngeal nerve in isolated gill preparations of trout for comparison (modified from Burleson & Milsom, 1993; see Discussion).

Results

Chronic hypoxia

5-HT-IR NECs of the gill filaments were organized in a linear pattern along the filament epithelium (Fig. 2A). As reported previously (Jonz & Nurse, 2003), the filament epithelium contains a population of 5-HT-negative NECs that are immunopositive for the synaptic vesicle marker SV2 (Fig. 2A; arrow). After in vivo exposure of zebrafish to chronic hypoxia (CHox), NEC morphology appeared to have changed, as indicated by an increase in size of 5-HT-IR cells (Fig. 2B) compared to normoxic controls (Nox). In addition, a greater number of SV2-IR 5-HT-negative NECs occupied the filament epithelium in a position adjacent to 5-HT-IR NECs in CHox zebrafish (Fig. 2B; arrows). Figure 2 illustrates another striking morphological change in 5-HT-IR NECs associated with CHox. In Nox controls, 5-HT-IR NECs occasionally extended neurone-like processes that associated with zn-12-IR nerve fibres of the intrinsic nerve bundle (iNB; Fig. 2C and D). In gill filaments of zebrafish exposed to CHox, these processes appeared to be longer and more prevalent, and terminated at zn-12-IR nerve fibres with 5-HT-IR ‘end feet’ (Fig. 2E and F).

Figure 2. Chronic hypoxia induced hypertrophy, proliferation and process extension in gill neuroepithelial cells (NECs)in vivo.

Figure 2

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Zebrafish were exposed to either normoxia (control; A, C and D) or chronic hypoxia (B, E and F). NECs were identified by serotonin (5-HT, green) or SV2 (red) immunoreactivity (IR) and nerve fibres were labelled with zn-12. A, in controls, 5-HT-IR and SV2-IR shown together indicated colocalization of both markers in most NECs (yellow) that were arranged in a linear pattern along the filament epithelium (FE), and exposed a population of SV2-IR 5-HT-negative NECs (arrow). Mean NEC area, 44.1 ± 1.6 µm2. Scale bar 20 µm. B, after exposure to chronic hypoxia, 5-HT/SV2-IR NECs of the FE were similar in number and arrangement to those of control in A, but had increased in size. Mean NEC area 59.6 ± 2.4 µm2. In addition, the number of SV2-IR 5-HT-negative NECs (arrows) increased in comparison to normoxic controls. Same magnification as in A. C, 5-HT-IR NECs of the filament epithelium occasionally extended small processes (arrow) in control zebrafish. Scale bar 10 µm. D, same image as in C showing also the close association of NECs with a zn-12-IR intrinsic nerve bundle (iNB). E, in zebrafish exposed to chronic hypoxia, 5-HT-IR NECs extended long neurone-like processes that often terminated with ‘end feet’ (arrows). F, same image as in E showing that process end feet terminated at a zn-12-IR intrinsic nerve bundle (iNB).

Morphometric analysis of 5-HT-IR gill NECs revealed several significant differences between fish exposed to CHox (n = 33) and Nox controls (n = 25). As suggested from morphological observations, the mean projection area of 5-HT-IR NECs in CHox fish was 59.4 ± 1.8 µm2 (mean ± s.e.m), a value significantly greater (P < 0.005) than that of Nox controls (51.7 ± 1.6 µm2), corresponding to an increase in size of approximately 15% (Fig. 3A). Analysis also indicated significant changes in aspect ratio (Nox, 1.6 ± 0.03; CHox, 1.9 ± 0.06; P < 0.001) and shape factor (Nox, 0.75 ± 0.02; CHox, 0.70 ± 0.03; P < 0.05), indicating that NECs of CHox fish were longer and more complex in shape than those of controls. These changes were suggestive of an increase in surface area or process extension induced by CHox. Indeed, significantly more (P < 0.001) NECs extended cytoplasmic, neurone-like processes toward zn-12-IR nerve fibres in CHox zebrafish compared to Nox controls (Fig. 3B). In addition, NEC processes in CHox animals were significantly longer (P < 0.001) than those of controls (Fig. 3C). Finally, while the number of NECs with detectable amounts of 5-HT did not change, significantly more (P < 0.05) SV2-IR NECs were present in gill filaments of CHox zebrafish compared to Nox controls (Fig. 3D). As SV2-IR NECs included both 5-HT-positive and 5-HT-negative NECs, these data indicate that there was an increase in the number of 5-HT-negative NECs after CHox.

Figure 3. Morphometric analysis of neuroepithelial cells (NECs) in gill whole-mount preparations of zebrafish exposed to normoxia or chronic hypoxia.

Figure 3

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NECs identified by serotonin (5-HT) or SV2 immunoreactivity (IR) were identified and sampled from the distal region of a single filament in fish exposed to normoxia (Nox, n = 25) or chronic hypoxia (CHox, n = 33). A, projection area (µm2) of 5-HT-IR NECs in the gills of zebrafish after exposure. In zebrafish exposed to CHox, NECs were significantly larger than those of controls (t test, *P < 0.005). B, number of 5-HT-IR NECs with processes. The number of 5-HT-IR NECs in the sampled area with neurone-like processes was significantly higher in zebrafish exposed to CHox (t test, *P < 0.001). C, length (µm) of 5-HT-IR NEC processes. Processes of 5-HT-IR NEC from CHox zebrafish were significantly longer than those of normoxic controls (t test, *P < 0.001). D, number of 5-HT-IR (filled bars) and SV2-IR NECs (hatched bars). 5-HT-IR NECs did not significantly increase in number after exposure to CHox (ANOVA-Bonferroni, P > 0.05). However, significantly more SV2-IR NECs were present in gill filaments of CHox zebrafish (n = 23) compared to controls (n = 15, ANOVA-Bonferroni, *P < 0.05), indicating an increase in the number of NECs not immunoreactive for 5-HT. Values are mean ± s.e.m.

Morphological and passive membrane properties of isolated NECs

NECs of the zebrafish gill were identified in vitro for electrophysiological experiments using the vital dye Neutral Red (NR; Stuart et al. 1974; Youngson et al. 1993). Compartmentalized uptake of the dye, presumably into cytoplasmic dense-cored granules, typically characterized the staining pattern of NR (Fig. 4A, arrow). Immunocytochemical procedures verified that many NR-positive cells were also 5-HT-IR (Fig. 4A–D). The distribution of the diameters of isolated 5-HT-IR cells (n = 372) was bimodal, and included smaller NECs of the respiratory lamellae and larger NECs of the filaments, with modes at ∼5.1 µm and ∼6.9 µm, and an overall mean of 5.7 ± 0.04 µm (Fig. 4E). As the larger NECs of the gill filaments were sensitive to the effects of CHox (Fig. 3), and permitted more stable recordings, these were selected for electrophysiological experiments. Whole-cell, voltage- and current-clamp recordings were obtained from the larger NR-positive NECs of the gill filaments 24–48 h after isolation. Under voltage clamp, the mean membrane capacitance (Cm) of these cells was 4.9 ± 0.2 pF (n = 35), and input resistance (Rin; see later) was 1.3 ± 0.3 GΩ (n = 10). The resting membrane potential (Vm) measured under current clamp (I = 0 pA) was −52.5 ± 2.1 mV (n = 15).

Figure 4. Isolation of neuroepithelial cells (NECs) of zebrafish gill filaments.

Figure 4

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A, bright field image of a NEC (arrow) stained with Neutral Red (NR) 24 h after enzymatic dissociation. Note the typical compartmentalized staining pattern of NR. Other cells were not stained with NR (arrowheads). Scale bar 10 µm. B, phase contrast image of cells in A after fixation. C, the fluorescence image shows that only the NR-positive NEC shown in A and B is immunoreactive for serotonin (5-HT, green). D, images in B and C merged together. E, frequency distribution of the diameter of dissociated gill NECs. Measurement of the diameter of dissociated 5-HT-immunoreactive NECs (n = 372) indicated the presence of two populations, as evidenced by modes at ∼5.1 µm and ∼6.9 µm (dashed lines). Larger NECs of the gill filaments were selected for patch-clamp recording.

NECs express an O2-sensitive outward current

Approximately 60% of NECs (i.e. 41 out of 68) investigated under voltage or current clamp responded to hypoxia (PO2 = 25 mmHg). Under voltage clamp, NECs expressed an outwardly rectifying current that was sensitive to changes in PO2. The inhibitory effects of hypoxia ranged from ∼10% to 48% (n = 41) with an average of ∼16% (see below). Figure 5A depicts the I–V relationship obtained by a ramp protocol from a single cell in which outward current was reversibly inhibited by hypoxia over a broad range of membrane potentials (−40 mV to +50 mV). A plot of the I–V relationship of the mean response of 10 cells to hypoxia indicated a small reduction in inward leak current at potentials more negative than −80 mV and a decrease in outward current at −50 mV, a potential near resting Vm (Fig. 5A, inset). Membrane currents were unaffected in control experiments, where the extracellular perfusate was bubbled with compressed air instead of N2 (n = 4; not shown). The mean (n = 10) O2-sensitive difference current was obtained by subtracting the current evoked under hypoxia from that under normoxia (Fig. 5B) using the same ramp protocol as in Fig. 5A. An I–V plot revealed that this O2-sensitive current displayed open rectification, as described by the Goldman–Hodgkin–Katz current equation (dotted curve; Fig. 5B). The mean reversal potential (Vr) was −75.8 ± 2.0 mV (n = 10), a value near the Nernst potential for K+ (EK), calculated to be −83 mV. These data suggest that the O2-sensitive current (IKO2) is carried by background K+-selective ion channels. Estimation of Rin from the slope of the I–V traces between −50 mV and −70 mV revealed that hypoxia significantly increased Rin from 1.3 ± 0.3 to 1.7 ± 0.4 GΩ (n = 10, paired t test, P < 0.01). Thus, hypoxia decreased K+ conductance in NECs near the resting membrane potential.

The degree of inhibition of IK by hypoxia was dependent upon O2 tension. The time course of IK inhibition in NECs (n = 8) during bath perfusion with hypoxic solution is shown in Fig. 5C, where currents were evoked by voltage steps from −60 mV to +30 mV every 10 s. The PO2 of the perfusate in the recording chamber was monitored with a carbon fibre electrode under identical experimental conditions. IK decreased progressively as the PO2 fell from ∼140 mmHg to ∼25 mmHg. As previously shown by Montoro et al. (1996), direct monitoring of O2 tension in the recording chamber during ramp-like application of hypoxia allows observation of the O2-dependence of IK. Voltage-clamp and PO2 data from Fig. 5C were used to generate the percentage inhibition of IK as a function of PO2 (Fig. 5D). Mean (n = 8) IK inhibition was 16.6 ± 2.5% at 25 mmHg.

Pharmacological characterization of IKO2

Voltage-clamp experiments were performed in which the K+ channel blockers, TEA, 4-AP and quinidine were used to pharmacologically characterize IKO2. To determine whether IKO2 was carried by voltage-dependent K+ channels, TEA and 4-AP were used in combination. As shown in Fig. 6A and B, O2-sensitive NECs were first identified by the occurrence of hypoxic inhibition of outward current. In such cells, addition of 20 mm TEA plus 5 mm 4-AP to the extracellular perfusate caused inhibition of the outward current by 19.8 ± 8.1% (n = 5), during a voltage step to +30 mV. However, hypoxic inhibition of outward current still occurred in the presence of TEA and 4-AP, suggesting that IKO2 was not carried by voltage-dependent K+ channels (Fig. 6A and B). As shown for a group of five cells in Fig. 6C, IKO2 current density (pA pF−1), calculated at test potentials of 0 mV and +30 mV, was not significantly different in the absence (control) and presence of TEA and 4-AP (P > 0.05). However, it should be noted that the intracellular Ca2+ concentration was clamped at relatively low levels (< 1 nm), and may not have been adequate for the detection of Ca2+-dependent K+ channels.

Figure 6. IKO2was insensitive to tetraethylammonium (TEA) and 4-aminopyridine (4-AP).

Figure 6

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A, typical whole-cell recording from an O2-sensitive NEC showing the persistence of the effects of hypoxia in the presence of 20 mm TEA and 5 mm 4-AP. The cell was held at −60 mV and stepped to +30 mV; current traces are shown for control (Cont), hypoxia (Hox) TEA + 4-AP, and TEA + 4-AP + Hox. B, I–V relationship from another O2-sensitive cell similar to A, after steps to a range of potentials. Application of 20 mm TEA and 5 mm 4-AP (•) reduced K+ current relative to control (Cont, ○). Hypoxic inhibition of residual K+ current in the presence of TEA and 4-AP (▪) was observed over a range of test potentials; effects of hypoxia alone omitted for clarity. C, mean ± s.e.m. (n = 5) current density (pA pF−1) of IKO2 (i.e. Nox–Hox) recorded at 0 mV and +30 mV test potentials under control conditions (filled bars, a–b in A) and in the presence of 20 mm TEA and 5 mm 4-AP (hatched bars, c–d in A). IKO2 was not significantly different in the presence of TEA and 4-AP compared to control at either test potential (paired t test, P > 0.05). Recovery from the effects of TEA plus 4-AP was ∼96% of initial control values at +50 mV (n = 5).

In order to confirm that background K+ channels were the likely mediators of O2-sensitivity in NECs, the effect of 1 mm quinidine, a non-specific blocker of these channels (O'Kelly et al. 1999; Buckler et al. 2000; Campanucci et al. 2003), was tested. The effects of hypoxia on outward current in a single cell in the absence (control) and presence of 1 mm quinidine is shown for step depolarizations to +30 mV (Fig. 7A) and ramp depolarizations (Fig. 7B). In this O2-sensitive NEC, quinidine caused a marked inhibition of outward current over a wide range of test potentials (93.5 ± 3.0% at +30 mV, n = 5), and moreover, the presence of quinidine occluded the hypoxic response (Fig. 7A and B). The I–V relationship of the O2-sensitive difference current IKO2 in control conditions and in the presence of 1 mm quinidine is shown in Fig. 7C. For a group of five cells treated in this way, the IKO2 current density (pA pF−1) in the absence (control) and presence of quinidine at −50 mV (i.e. near resting Vm for NECs) and 0 mV is shown in Fig. 7D. Quinidine reduced IKO2 from 0.94 ± 0.37 to −0.16 ± 0.05 pA pF−1 at −50 mV (paired t test, P < 0.05), and from 3.4 ± 1.1 to −0.6 ± 0.05 pA pF−1 at 0 mV (paired t test, P < 0.005), indicating blockade of most or all of the hypoxic response. This marked sensitivity of IKO2 to quinidine, together with its insensitivity to TEA and 4-AP (Fig. 6), is consistent with the presence of O2-sensitive background K+ channels in NECs.

Figure 7. Effects of quinidine onIKO2.

Figure 7

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A, typical whole-cell recording from an O2-sensitive NEC showing the occlusion of the hypoxia response by 1 mm quinidine (Quid). The cell was held at −60 mV and was stepped to +30 mV. The application of hypoxia (Hox) reduced IK compared to control (Cont). However, hypoxic inhibition of IK was not evident when the experiment was performed in the presence of quinidine (Quid + Hox). B, I–V relationship obtained using a ramp protocol from the cell shown in A. Hypoxia (Hox) reduced IK over a range of potentials, but had little effect in the presence of 1 mm quinidine (Quid + Hox). C, average I–V relationship of IKO2 from five cells in control (Cont, a–b in A and B) and in the presence of 1 mm quinidine (Quid, c–d in A and B). D, mean ± s.e.m. (n = 5) current density (pA pF−1) of IKO2 (i.e. Nox–Hox) recorded at −50 mV and 0 mV test potentials under control conditions (Cont, a–b in A and B) and in the presence of 1 mm quinidine (Quid, c–d in A and B). IKO2 was significantly reduced or abolished (asterisks) at both test potentials in the presence of quinidine (paired t test, P < 0.05). Recovery from the effects quinidine was ∼75% of initial control values at +50 mV (n = 5).

Effects of hypoxia and K+ channel blockers on membrane potential

Experiments described above suggest that hypoxia inhibits background K+ channels that are open at the resting potential of NECs. If so, we predicted that hypoxia should depolarize NECs, resulting in a receptor potential under current-clamp conditions. As expected, hypoxia caused a reversible depolarization associated with a conductance decrease, or an increase in Rin, in NECs (Fig. 8A). In five cells, there was a significant decrease in Vm from −50.6 ± 3.1 to −44.2 ± 3.4 mV (paired t test, P < 0.005) after bath perfusion with hypoxic solution (Fig. 8B), corresponding to a depolarization of ∼6 mV. As shown in Fig. 8F, hypoxia caused a significant increase in Rin from 1.7 ± 0.6 to 2.4 ± 0.7 GΩ (n = 5; paired t test, P < 0.05).

Figure 8. Effects of hypoxia and quinidine on membrane potential (Vm) and input resistance (Rin) in isolated neuroepithelial cells (NECs).

Figure 8

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A, typical current-clamp recording of a reversible depolarization of Vm during perfusion (bar) with hypoxic solution (Hox; PO2 = 25 mmHg); resting potential (Vm) was ∼ −50 mV. Injection of 10 pA of hyperpolarizing current for 1 s (downward deflections) evoked a greater change in voltage during hypoxia, indicative of an increase in Rin. B, mean ± s.e.m. (n = 8) Vm of NECs in normoxia (Nox) versus hypoxia (Hox); Vm was significantly, and reversibly, reduced by ∼6 mV in the presence of Hox relative to Nox controls (paired t test, *P < 0.005). C, typical current-clamp recording showing the pronounced depolarization due to application of 1 mm quinidine (Quid) on a cell that was initially at a resting potential of −53 mV; perfusion with hypoxic solution had no effect on Vm in the presence of 1 mm Quid. Dashed line indicates a 6-min pause in recording during recovery. D, current-clamp recording of hypoxic depolarization following administration of 20 mm TEA and 5 mm 4-AP. TEA and 4-AP alone did not change Vm. E, effect of 1 mm quinidine (Quid) on hypoxia-induced depolarization. Mean ± s.e.m. (n = 5) change in membrane potential (ΔVm) in response to hypoxia (Hox) was significantly reduced or eliminated in the presence of Quid (paired t test, *P < 0.05). F, hypoxia-induced changes in Rin (n = 5) are occluded in the presence of quinidine (Quid). Rin was calculated using the equation Rin = ΔV ΔI−1 and the current injection protocol outlined in A. In control (Cont), a significantly greater Rin was observed after perfusion with hypoxic (Hox) solution compared to normoxic (Nox) solution (paired t test, *P < 0.05). The effects of 1 mm Quid increased Rin under normoxic conditions and prevented any further change after addition of the hypoxic perfusate (paired t test, P > 0.05).

Because quinidine blocked IKO2 in NECs, as demonstrated in voltage-clamp experiments (Fig. 7), it was predicted that this drug should also abolish the hypoxic depolarization. As exemplified in Fig. 8C, 1 mm quinidine produced a robust depolarization that caused Vm to change from −53 to +18 mV. This pronounced depolarization was not unlike that seen in O2-sensitive H-146 cells in response to quinidine (O'Kelly et al. 1999). Moreover, application of quinidine to the recording chamber occluded the effects of hypoxia on Vm (Fig. 8C). Data from five cells treated in this way are summarized in Fig. 8E. The hypoxia-induced depolarization (5.9 ± 0.9 mV) was significantly reduced in the presence of 1 mm quinidine (0.1 ± 1.2 mV; paired t test, P < 0.05). Similarly, Fig. 8F shows that quinidine produced a corresponding increase in Rin (5.5 ± 2.3 GΩ) that did not change significantly after perfusion with hypoxic solution (5.5 ± 1.8 GΩ, paired t test, P > 0.05). The effects of quinidine on Vm and Rin were fully reversible upon washout of the drug. In contrast to the effects of quinidine, addition of 20 mm TEA and 5 mm 4-AP to the bath had no effect on Vm (n = 2), and the hypoxia-induced depolarization (> 5 mV) persisted in the presence of these drugs (Fig. 8D). Together, these data show that membrane depolarization and increases in Rin during hypoxia are mediated by inhibition of quinidine-sensitive K+ channels.

Discussion

Morphological changes induced by chronic hypoxia

The combination of confocal immunofluorescence and morphometric analysis revealed several changes in NEC morphology following in vivo exposure to chronic (60 days) hypoxia (CHox). The projection area of serotonin (5-HT)-immunoreactive (IR) NECs, which is a measure of cell size, increased by approximately 15%, and this was associated with an increase in aspect ratio (AR) and a decrease in shape factor (SF). An increase in AR is indicative of cell elongation, while a decrease in SF demonstrates an increase in cell surface area and complexity of shape, as observed during neuronal growth, differentiation and neurite extension (Levallois et al. 1997). Further analysis revealed that CHox had induced the growth and extension of 5-HT-IR neurone-like processes from NECs that made contact with fibres of the intrinsic nerve bundle (iNB) of the gill filament. NEC processes in CHox zebrafish contained 5-HT and terminated with enlarged ‘end feet’, which may act to increase surface area and enhance neurotransmitter release onto postsynaptic fibres. These changes demonstrate that growth and plasticity of NECs may be regulated by O2 tension, and further implicate the iNB as an important pathway involved in the hypoxia response (see Fig. 1B), as suggested in a previous study (Jonz & Nurse, 2003).

In addition to the morphological changes just described, exposure to CHox induced proliferation of NECs within the filament epithelium. However, only 5-HT-negative NECs, labelled with the synaptic vesicle marker, SV2 increased in number. This suggests that they too may play a role in O2 sensing. It is surprising that the number of NECs containing detectable amounts of 5-HT did not appear to change after CHox. In light of the ultrastructural observation that NECs of the fish gill appeared to degranulate (suggestive of exocytosis) after exposure to acute hypoxia (Dunel-Erb et al. 1982), and that in functional studies hypoxia induced 5-HT release in neuroepithelial bodies (NEBs) of the mammalian lung (Cutz et al. 1993; Fu et al. 2002), it is plausible that in our studies depletion of 5-HT after exocytosis may have masked our ability to detect the full extent of NEC proliferation after CHox. Alternatively, 5-HT-negative NECs in zebrafish may be an immature form of NECs still undergoing differentiation (Jonz & Nurse, 2003). Proliferation of these cells during acclimatization to hypoxia may act to increase cell renewal and ultimately increase the number of NECs expressing 5-HT. Nevertheless, it is evident that the density of NECs within the filament epithelium is regulated by O2 tension.

The present study demonstrates the plasticity and remodelling of the NEC–nerve fibre system of the zebrafish gill filaments following exposure to CHox, and suggests that these O2-regulated changes may have physiological significance during acclimatization to hypoxia. Increases in both ventilatory frequency (Kerstens et al. 1979; Johnston et al. 1983) and amplitude (Burleson et al. 2002) have been demonstrated in fish following acclimation (1–4 weeks) to hypoxia. Similar changes in the morphology and number of O2-sensitive type I cells of the mammalian carotid body are associated with exposure to CHox. Chemoreceptor hypertrophy and proliferation occurred after in vitro (Stea et al. 1992; Mills & Nurse, 1993; Nurse & Vollmer, 1997) and in vivo (for review see Wang & Bisgard, 2002) exposure to hypoxia. In addition, CHox induced changes in ion channel expression (Stea et al. 1992; Wyatt et al. 1995) and neurotransmitter levels (Jackson & Nurse, 1997; Bisgard, 2000) in type I cells, and altered sensitivity of the carotid body to hypoxia (González et al. 1994; Bisgard, 2000). Such changes may be central to the importance of peripheral O2 chemoreceptors in maintaining ventilatory drive during acclimatization to hypoxia in mammals (Forster et al. 1981; Bisgard, 2000) and in other vertebrates, such as fish.

Electrophysiological characterization of O2 sensitivity

Whole-cell voltage- and current-clamp recordings were performed on isolated NECs, permitting electrophysiological characterization of a cellular response to hypoxia. NECs immunoreactive for 5-HT were present in culture, and these were identified in vitro using the vital dye Neutral Red (Stuart et al. 1974; Youngson et al. 1993). Hypoxia produced a dose-dependent, reversible inhibition of outward whole-cell current in NECs. The I–V relationship of the O2-sensitive difference current (IKO2) reversed near the equilibrium potential for K+ (EK = −83 mV) and displayed open rectification, as described by the Goldman–Hodgkin–Katz current equation. Such characteristics strongly suggest that IKO2 was carried by K+ ions through background or ‘leak’ K+ channels, which are open at resting membrane potentials (Goldstein et al. 2001; see also Buckler, 1997). Furthermore, hypoxia increased input resistance (Rin) near the resting membrane potential, consistent with the hypothesis of a hypoxia-induced inhibition of background K+ channels.

Pharmacological characterization of IKO2 in NECs was consistent with the presence of O2-sensitive, background K+ channels. The O2-sensitive current was resistant to TEA and 4-AP, classical blockers of voltage-dependent, delayed rectifier K+ currents. However, IKO2 was virtually abolished in the presence of 1 mm quinidine across a range of test potentials. Similarly, O2-sensitive background K+ currents identified in other chemoreceptor cells, such as type I cells of the carotid body (Buckler, 1997; Buckler et al. 2000), peripheral neurones (Campanucci et al. 2003), and NEB-derived H-146 cells (O'Kelly et al. 1999), were resistant to TEA and 4-AP, but displayed strong sensitivity to 1 mm quinidine. Thus, the present biophysical and pharmacological data suggest the involvement of a background K+ conductance in mediating the O2 sensitivity of NECs of the gill.

Current-clamp experiments demonstrated that NECs responded to hypoxia with a reversible depolarization of ∼6 mV. In accordance with voltage-clamp data, this hypoxia-induced receptor potential persisted in the presence of TEA and 4-AP, but was abolished following incubation with quinidine, which itself, caused a robust depolarization of NEC membrane potential. In light of evidence from other neuroepithelial sensory receptors that express voltage-dependent Ca2+ channels, such as taste cells and Merkel cells (Kinnamon & Roper, 1988; Yamashita et al. 1992), we propose that depolarization of gill NECs during exposure to hypoxia leads to Ca2+ entry via voltage-dependent Ca2+ channels and neurosecretion (possibly 5-HT) onto sensory afferent terminals, as described in the mammalian carotid body (González et al. 1994; López-Barneo et al. 2001). However, the nature of Ca2+ channels in zebrafish gill NECs awaits further study. In addition, further experimentation is required to determine the specific type of background K+ channel involved in O2 sensing by gill NECs (see Goldstein et al. 2001; Lesage, 2003). O2 sensing in carotid body type I cells, and in glossopharyngeal neurones that innervate the carotid body, appears to be mediated at least in part by background K+ channels similar to those of the acid-sensitive (TASK) and halothane-inhibited (THIK) families, respectively (Buckler et al. 2000; Campanucci et al. 2003). In both cases, hypoxic modulation of channel activity was dependent upon the presence of intracellular factors (Buckler et al. 2000; Campanucci et al. 2003). However, in the present study, O2-sensitive NECs were dialysed during whole-cell recordings, suggesting that K+ channel inhibition occurred in the absence of cytoplasmic modulators (see also Kemp et al. 2004). Nevertheless, this study raises the possibility that during the evolution of O2 sensing in vertebrates, hypoxic regulation of voltage-dependent K+ channels in peripheral chemoreceptors arose as a later event, perhaps in air-breathers, whereas regulation of voltage-independent or background K+ channels by hypoxia has been relatively conserved.

Physiological significance of gill NECs

This study has characterized the O2 sensitivity of NECs of the zebrafish gill and strongly supports a role for these cells as hypoxic chemoreceptors, analogous to those of the mammalian carotid body (González et al. 1994) and pulmonary NEBs (Youngson et al. 1993; Fu et al. 2002). Several studies have indicated that, in fish, the hypoxic response appears to be initiated by two types of O2 chemoreceptors in the gill. One type is internally orientated, detecting changes in arterial PO2, and the other is externally orientated, monitoring environmental changes in PO2 (for review see Burleson et al. 1992). Although NECs of the gill filaments in zebrafish were not observed to be directly exposed to the external environment (Jonz & Nurse, 2003), they are ideally situated within a permeable epithelium between the respiratory water flow and the arterial blood supply (Fig. 1A and B; Jonz & Nurse, 2003) and may be capable of sensing changes in PO2 from both environments without direct exposure (Perry & Gilmour, 2002). As shown in Fig. 5D, our data indicate that the response curve (i.e. IK inhibition) of zebrafish NECs to changes in PO2 closely resembles the response (i.e. increase in discharge frequency) to hypoxia recorded from sensory fibres innervating the gill arch in isolated gill preparations of trout (Burleson & Milsom, 1993). This suggests that the chemoreceptor response in the gill during hypoxia may arise from activation of sensory pathways initiated by IK inhibition and depolarization in NECs. However, this comparison between IK inhibition and nerve discharge is limited by the absence of available information on Ca2+ channels and neurosecretion in NECs. In addition, the evolutionary relationship between gill NECs and carotid body type I cells suggests that NECs may act as O2 chemoreceptors. The mammalian carotid body and the first gill arch in fish are derived from the same embryonic pharyngeal arch (Weichert, 1967) and both receive innervation from the glossopharyngeal and vagus nerves (Nilsson, 1984; González et al. 1994). However, as NECs are invariably present on all gill arches in zebrafish (Jonz & Nurse, 2003), it appears that O2 sensing is not limited to the first gill arch.

Results from this and a previous study (Jonz & Nurse, 2003) suggest that stimulation of NECs of the zebrafish gill by low PO2 may lead to activation of local and centrally mediated sensory pathways in response to hypoxia (see also Fig. 1B). Activation of these pathways during hypoxia may occur by secretion of neurotransmitters contained within synaptic vesicles, which are located in the basal cytoplasm of NECs (Jonz & Nurse, 2003). Although identification of the chemical transmitter(s) involved in hypoxic chemotransmission is premature, evidence from this and another study (Dunel-Erb et al. 1982) employing exposure to hypoxia may indicate the involvement of 5-HT. In fish, exogenous application of a number of neurochemicals, including 5-HT, leads to an increase in nerve discharge in isolated gill preparations (Burleson & Milsom, 1995a) and to changes in gill vascular resistance, ventilation and heart rate (Burleson et al. 1992; Fritsche et al. 1992; Burleson & Milsom, 1995b; Sundin et al. 1995, 1998).

We have defined a novel preparation for studying the cellular mechanisms of O2 chemoreception in aquatic vertebrates, which allows electrophysiological investigation of isolated O2-sensitive cells combined with detailed morphological examination of these cells in whole-mount tissue specimens. The zebrafish is tractable to large-scale genetic screens and has become an ideal vertebrate animal model amenable to studies of nervous system development (Eisen, 1996; Fishman, 1999) and the structure and function of sensory systems (Brockerhoff et al. 1998; Starr et al. 2004; Söllner et al. 2004). With the aid of genetic or developmental manipulations that may modify or perturb O2 sensing, the zebrafish may represent an attractive model for studying the subcellular mechanisms and development of O2 chemoreception, and the role of peripheral O2 chemoreceptors in respiratory regulation and acclimatization to hypoxia.

Acknowledgments

The authors thank Cathy Vollmer, Verónica Campanucci, Dr C. M. Wood, Dr M. J. O'Donnell and Dr M. Zhang for their comments during these studies. We also acknowledge the Natural Sciences and Engineering Research Council (NSERC) of Canada for funding through an operating grant to C.A.N., and NSERC and the Ontario Graduate Scholarship Program for postgraduate scholarships to M.G.J. Antisera, zn-12 and SV2, were obtained from the Developmental Studies Hybridoma Bank developed under the auspices of the NICHD and maintained by the University of Iowa, Department of Biological Sciences, Iowa City, IA, USA.

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