Molecular Vision 2007; 13:2163-2182 <http://www.molvis.org/molvis/v13/a247/>
Received 16 April 2007 | Accepted 29 October 2007 | Published 27 November 2007		 Download
Reprint

Voltage-gated sodium channel α-subunits Nav1.1, Nav1.2, and Nav1.6 in the distal mammalian retina

Deb K. Mojumder1, Laura J. Frishman1, Deborah C. Otteson1, David M. Sherry1,2
 
 

1University of Houston College of Optometry; 2University of Houston, College of Pharmacy, Department of Pharmacological and Pharmaceutical Sciences, Houston, TX

Correspondence to: Laura J. Frishman, University of Houston, College of Optometry, 505 J. Davis Armistead Bldg., Houston, TX 77204-2020; Phone: (713) 743-1972; FAX: (713) 743-2595 email: lfrishman@uh.edu
 
Dr. Mojumder is at the Verna and Marrs McLean Department of Biochemistry and Molecular Biology, Room N430, Baylor College of Medicine, Houston, TX, and Dr. Sherry is at University of Oklahoma Health Sciences Center, Department of Cell Biology, 940 Stanton L. Young Blvd, BMSB-536, Oklahoma City, OK

Abstract

Purpose: Recent studies indicate the presence of functional voltage-gated sodium channels (Nav channels) in the distal retina in several species. This study examined the distribution of Nav channels in the outer plexiform layer (OPL) of rat, mouse, and rabbit retinas.

Methods: Immunohistochemical and electroretinographic approaches were used.

Results: Antibodies specific for Nav1 α-subunits appropriately labeled retinal ganglion cells, their axons, and amacrine cells that are known to have tetrodotoxin (TTX)-sensitive Nav channels. Pan-Nav, Nav1.2, and Nav1.6 labeling was found in horizontal cells and processes in all three species. Weaker Nav1.1 labeling was observed in rodent horizontal cells, but some rabbit horizontal cells and processes were prominently labeled. Additional labeling for Nav1.1, Nav1.2, and Nav1.6 that was not attributable to horizontal cells was also present in the OPL. Much of this labeling was diffusely distributed. Some of the additional Nav1.1 labeling was associated with photoreceptor terminals. By exclusion using photoreceptor and horizontal cell markers, some of this labeling could have been associated with bipolar cell dendrites, although colocalization was not directly established due to the diffuse nature of the labeling and limits on anatomical resolution. No Nav1 α-subunit labeling was observed in bipolar cell bodies. Testing for functional Nav channels was performed by recording full field flash electroretinograms from dark-adapted rats before and after intravitreal injections of TTX, 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX), or TTX+CNQX. TTX and CNQX+TTX, but not CNQX alone, greatly attenuated the dark-adapted cone-driven b-waves.

Conclusions: Horizontal cells from three different mammalian retinas showed prominent labeling for Nav1 α-subunits. Some additional diffuse Nav1 α-subunit labeling in the OPL was associated with photoreceptor terminals. Nav1 α-subunit labeling also may have been present on bipolar cell dendrites, although it was not possible to establish this localization unequivocally by immunostaining. However, cone-driven b-waves in rats were reduced in maximum amplitude by TTX in the presence of CNQX which blocks synaptic input to horizontal, amacrine, and ganglion cells. This finding is consistent with TTX effects on the b-wave being due to blockade of Nav channels in cone bipolar cell dendrites in the OPL. The role of Nav channels in horizontal cells remains to be determined.

Introduction

Voltage-gated Na+ channels (Nav channels) are members of a large superfamily of ion channel proteins in membranes of cells that are characteristically involved in electrical signaling [1,2]. The opening of these channels in response to membrane depolarization causes an influx of Na+ ions to the cell that is essential for initiating and propagating sodium-dependent action potentials in neurons and muscle cells [3]. However, the functional role of Nav channels is not restricted to mediating rapid changes in membrane polarization and action potentials. Recent studies demonstrate that Nav channels have wider physiological and developmental roles. For example, Nav channels can load cells with Na+ ions during slow inactivation and thereby modulate intracellular Na+ homeostasis [4]. Nav channel interactions with tenascin and receptor protein tyrosine phosphatase-suggest these channels also can participate in cell-cell or cell-matrix interactions (for review see [5]). The absence of a specific Nav α-subunit (Nav1.6) during development leads to deficits in photoreceptor function as evidenced by reduced amplitudes of the electroretinogram (ERG) waves originating from photoreceptors as well retinal postreceptoral neurons [6].

Nav channels are composed of a large (about 260 kDa) pore-forming α-subunit (Figure 1A) associated with one or more smaller α-subunits [1,2,7]. The Nav α-subunit is obligatory for channel function [1,2,5]. There are ten known Nav α-subunit isoforms (designated Nav1.1 through Nav1.9 and Nax), each encoded by a separate gene [1,8,9]; (gene nomenclature is provided in Table 1). Nav1.1, Nav1.2, Nav1.3, and Nav1.6 are highly expressed in the central nervous system, including the retina [8-15]. These Nav1 α-subunits are all sensitive to tetrodotoxin (TTX) blockade, a property that has been used to demonstrate the presence of functional Nav channels in various retinal neurons (summarized in Figure 1B). The TTX sensitivity of retinal ganglion cell action potentials is well known [16-18]. Several subtypes of amacrine cells, the other major third order retinal neurons, also possess TTX-sensitive Na+ currents (e.g., AII, A17, A18, and starburst amacrine cells); [19-23].

TTX-sensitive Na+ currents also have been reported in other retinal neurons. These currents, however, are not as prominent as those of amacrine and ganglion cells, and their functional significance is not yet fully understood. Recent studies have identified voltage-gated, TTX-sensitive Na+ currents in classes of bipolar cells from a variety of vertebrate retinas: rat [24], goldfish [25], salamander [26], ground squirrel [27], and most recently, human [28,29]. In the goldfish, Na+ channel immunoreactivity was localized to cone bipolar cells of both ON and OFF types [25]. In rat retina, TTX-sensitive voltage-gated Na+ currents were also observed in subsets of ON and OFF cone bipolar cells, but not in rod bipolar cells. TTX-sensitive currents also have been detected in isolated retinal horizontal cells from several species, but have never been identified in intact retinal recordings: catfish [30], white perch [31], cat: A-type horizontal cells [32], rabbit: 81% of the A-type and 90% of the B-type cells [33], and mouse [34]. Voltage-gated Na+ channels also have been found in human rods and cones where their precise role is still unclear [35,36]. Together, these studies suggest a potentially broader role for Nav channels in the retina than has been appreciated to date.

Detailed knowledge of the distribution of the Nav α-subunits is important to interpreting the various functional roles of Nav channels and the different Nav α-subunits in physiology and other processes in the retina. To better understand Nav channels in the mammalian retina, we have examined the distribution of Nav channel α-subunits in the retina using immunohistochemical and ERG approaches, with particular focus on expression of Nav1.1, Nav1.2 and Nav1.6 by second-order neurons. To assess similarities and differences across different mammalian species, we performed experiments that used the retinas of rat, mouse, and, to a lesser extent, rabbit. Each Nav1 α-subunit showed a distinctive distribution in the retina. As expected, labeling was present in third-order retinal ganglion cells and amacrine cells. However, labeling for each Nav1 α-subunit also was found in horizontal cells and their processes and elsewhere in the outer plexiform layer (OPL). Application of TTX reduced the amplitude of the dark-adapted cone-driven b-wave of the ERG, an ERG component known to arise primarily from light-evoked responses of ON cone bipolar cells.

Methods

Animals for immunohistochemical studies

Studies were performed using nine adult Brown Norway rats (Rattus norvegicus, Charles River Laboratories, Wilmington, MA; six weeks-six months of age), six mice (Mus musculus, C57/BL6; Simonsen Laboratories, Gilroy, CA; six weeks-six months of age), and four rabbits (Oryctolagus cuniculus, New Zealand White; Myrtles Rabbitry Incorporated, Thompsons Station, TN; six to nine months of age). All animals were maintained on a 12 h: 12 h dark cycle with food and water available ad libitum. All animals were light-adapted at the time of euthanasia. Animals for immunohistochemical studies were anesthetized using an intramuscular injection of ketamine (Vedco Inc., St Joseph, MO; rats, 86 mg/kg; mice, 70 mg/kg; rabbits, 35 mg/kg, obtained from) and xylazine (Vedco Inc.; rats, 6.5 mg/kg; mice, 7 mg/kg; rabbits, 5 mg/kg) and then were euthanized by an intramuscular injection of a lethal dose of pentobarbital (Ft. Dodge Laboratories, Inc., Ft. Dodge, IA; 150-200 mg/kg). All animal procedures and protocols conformed to US Public Health Service and Institute for Laboratory Animal Research guidelines and were approved by the University of Houston Institutional Animal Care and Use Committee.

Tissue preparation for immunohistochemistry

To ensure that immunolabeling patterns obtained with each antibody were consistent and accurate we performed immunolabeling in multiple species and in two different tissue preparations. Tissue fixation regimes were optimized accordingly for each preparation and antibody. The effects of fixation and tissue preparation on labeling patterns is discussed further in Results.

Following euthanasia, the eyes used for frozen sections were rapidly excised from the head, leaving a portion of the superior rectus muscle to indicate the superior pole of the globe. The corneas were slit with a razor blade, the lens was removed, and eyes were immersed in 4% paraformaldehyde in 0.1 M cacodylate buffer (pH 7.4) for 15 min to 2 h at 4 °C. Following fixation, eyes were rinsed in phosphate buffered saline (PBS; pH 7.4) and cryoprotected overnight in 30% sucrose/PBS overnight. The next day, the eyes were embedded in Tissue-Tek OCT Compound (Tissue-Tek, Hatfield, PA) and fast frozen in liquid nitrogen. Vertical cryostat sections of 10-12 μm thickness were taken along the vertical meridian of the eyecup and collected onto Superfrost/Plus microscope slides (Fisherbrand, Fisher Scientific, Pittsburgh, PA). Sections were stored at -20 °C until use.

Retinal wholemounts were prepared by excising the eye and removing the cornea and lens as described in the previous paragraph The neural retina was rapidly isolated free of the retinal pigmented epithelium in cold Ames' medium (4 °C, pH 7.4) and then immediately fixed lightly in 4% paraformaldehyde in 0.1 M cacodylate buffer (pH 7.4) for 5 min at 4 °C. The vitreous humor was removed, and relaxing cuts were made in the retinal margin to allow the retina to flatten. The retina was rinsed in PBS and immunolabeled free-floating.

Antibodies and antisera

The details of all primary antibodies are presented in Table 2. Primary antibodies against Nav1 α-subunits were mouse monoclonal antibodies raised against relatively short amino acid epitopes based on rat Nav1 α-subunit sequences and have been used previously for studies of Nav1 α-subunits in the retina and optic nerve [13,14]. Figure 1A provides a schematic representation of the structure of the Nav1 α-subunit and shows the locations of the amino acid epitopes used to raise each Nav1 α-subunit antibody. The specificity of these antisera has been extensively characterized previously using immunoblotting, ELISA and preabsorption assays and immunolabeling of cells and tissues known to either contain or lack specific Nav1 α-subunits [13,14,37] (Dr. James Trimmer, personal communication). A pan-specific antibody that recognizes all Nav1 α-subunits (Pan-Nav) was raised against a synthetic peptide corresponding to a region of the intracellular III-IV loop conserved in all Nav1 family α-subunits [37] (Figure 1A for a schematic representation of the structure of the Nav1 α-subunit). Anti-Nav1.1 was raised against a fusion protein corresponding to amino acids 1929-2009 of the C-terminal domain of rat Nav1.1 [14]. Anti-Nav1.2 was raised against a fusion protein corresponding to amino acids 1882-2005 of the C-terminal domain of rat Nav1.2 (Dr. James Trimmer, personal communication). The antibody specific for Nav1.6 was raised against a synthetic peptide corresponding to amino acids 459-476 of the intracellular interdomain loop I-II of rat Nav1.6 [13]. Immunoblotting of crude membrane homogenates from mouse retina and brain as described previously [38] confirmed labeling of a single band of appropriate molecular mass by the Nav1 α-subunit antibodies. A dilution series for each Nav1 α-subunit antibody was performed to determine the optimal dilution for immunolabeling. The specificity of the Nav1 α-subunit antibodies and epitopes is assessed further in Results. A panel of additional primary antibodies and antisera directed against a number of well-characterized cell- and synapse-specific markers also were used (see Table 2).

Immunoblotting of crude membrane homogenates from mouse retina and brain was used to confirm labeling of a single band of appropriate mass by the Nav1 α-subunit antibodies. For immunoblotting, neural retinas were isolated, or brain removed, and crude membrane homogenates were prepared by sonication and centrifugation [38]. Proteins were loaded onto 10% polyacrylamide gels and resolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Proteins were then transferred to polyvinylidene difluoride (PVDF) membranes for visualization. PVDF membranes were blocked using 2% normal goat serum in 0.1 M tris-buffered saline+0.05% tween-20 (TBS-Tween, pH 7.2), and then incubated overnight in primary antibody against Nav1.1, Nav1.2, or Nav1.6 at 4 °C. Blots were rinsed in TBS-Tween followed by TBS alone and incubated for 2 h at room temperature in goat anti-mouse-horseradish peroxidase (1:2000). Blots were rinsed in TBS, and labeled protein bands were visualized using an enhanced chemiluminescence kit according to the manufacturer's instructions (Santa Cruz Biotechnology, Santa Cruz, CA). All antibodies were diluted in 2% normal goat serum in TBS-Tween.

Binding of primary antibodies was detected using fluorescent secondary antisera. Secondary antisera were raised in goat and were specific for mouse, rabbit or sheep, immunoglobulins and were conjugated to Cy3 or Cy5 (diluted 1:200; Jackson ImmunoResearch Laboratories, West Grove, PA) or conjugated to AlexaFluor488 or AlexaFluor 546 (diluted 1:200; Molecular Probes, Eugene, OR). Double labeling was performed with an appropriate combination of secondary antibodies conjugated to different fluorescent tags.

Immunolabeling

In most experiments, immunolabeling of frozen sections employed immunofluorescent methods [38-40]. Briefly, frozen sections were thawed, immersed in 4% formaldehyde for 10-15 min at room temperature to improve adherence, and rinsed in deionized water. They were then treated with 1-2% NaBH4 for 1-2 min at room temperature to reduce non-specific autofluorescence, rinsed in deionized water, and exchanged to PBS. Non-specific labeling was blocked with 10% normal goat serum + 5% bovine serum albumin + 0.5-1% fish gelatin + 0.1-0.5% Triton X-100 in PBS (blocker). Excess blocker was removed, and primary antibody was applied for 2 days at 4 °C. For double labeling experiments, a combination of primary antibodies was applied simultaneously. Sections were rinsed, blocked for 30 min at room temperature, and secondary antibody was applied for 1 h at room temperature. For double-labeling experiments, an appropriate combination of secondary antisera was applied simultaneously. Sections were rinsed and coverslipped in a fade-retardant mounting medium (either Vectashield, Vector Labs, Burlingame, CA, or Prolong Gold, Molecular Probes, Eugene, OR) and examined in the microscope. Sections processed substituting normal rabbit serum for rabbit polyclonal primary antisera, non-specific mouse IgGs, or in the absence of primary antibodies showed no labeling, as appropriate. All primary and secondary antibodies were diluted in blocker.

To obtain consistent immunolabeling in frozen sections using the Pan-Nav antibody, an antigen retrieval step was required [41]. For immunolabeling of frozen sections using the Pan-Nav antibody, thawed sections were heated to 90 °C in PBS at pH 7.4-8.3. They were allowed to cool to room temperature and then were processed as described in the previous paragraph for frozen sections, without any further exposure of the sections to fixatives (See Results for further discussion).

Rat retinal wholemounts were immunolabeled free-floating. Wholemounts were treated with 1-2% NaBH4 for 1-2 min, rinsed in deionized water followed by PBS and incubated in blocker solution for 1 h at room temperature to block non-specific labeling. Retinas were incubated in primary antibody for 5 days at 4 °C. They were then were rinsed in PBS for 2 h at room temperature and incubated free-floating in secondary antibody at room temperature for 1 h. Retinas were rinsed in PBS for 2 h at room temperature, flattened onto microscope slides with the ganglion cell side up, coverslipped with Prolong Gold and examined in the confocal microscope.

Imaging

Images were acquired using a Leica TCS SP2 confocal microscope and LCS software (Leica Microsystems, Exton, PA). Images were captured using 20x (NA, 0.7), 63x oil (NA, 1.32), or 63x water immersion (NA, 1.2) objective lenses. Stacks of serial optical sections ranging from 1.5- 6 μm thickness in the Z-plane were collected using step sizes of 0.15-0.6 μm thickness. Assessment of labeling in single optical planes was performed using 63x objectives to achieve a maximal Z-plane resolution of approximately 0.3 μm. Images in each fluorescent channel were collected sequentially with laser power and detector sensitivity adjusted to prevent bleedthrough of signals between fluorescence channels. The absence of bleedthrough between channels was confirmed in sections treated with a single primary antibody and a combination of secondary antibodies to verify that only the channel corresponding to the primary antibody showed labeling.

Figures were prepared using Adobe Photoshop 6.0 (Adobe Systems, Inc., Mountain View, CA). Image scale was calibrated and image brightness and contrast were adjusted to highlight specific labeling, if needed.

Bioinformatics and sequence analysis

Sequences of orthologous Nav1 α-subunit proteins from Rattus norvegicus (rat), Mus musculus (mouse), Canis familiaris (dog), Bos taurus (cow), and Homo sapiens (human) were obtained from the NCBI protein database. Amino acid sequences for rat-specific peptides used as immunogens to generate the monoclonal antibodies against Nav1.1, Nav1.2 and Nav1.6 were obtained from the NeuroMab (Davis, CA). Protein sequences for Oryctolagus cuniculus (rabbit) were obtained by employing full length rat Nav1.x cDNA or protein sequences as queries for tBlastn or tBlastx searches of the translations of the predicted transcripts (Ab-initio cDNAs/Genscan/SNAP) in the Ensembl rabbit Genome Database (v42.1-Dec2006). Amino acid sequences were aligned using AlignX software (Vector NTI Advance 10.1; Invitrogen Corporation, Carlsbad, CA), using the rat peptides as the consensus sequence. For proteins within a species that were represented by more than one sequence in the database, multiple protein sequences were aligned to determine the consensus sequence. Rat peptide sequences were also used as a query for Basic Local Alignment Search Tool for Proteins (BlastP) and tBlastn searches against the protein, non-redundant and expressed sequence tag (EST) databases (NCBI), limiting search to mammalian sequences [42].

Electroretinograms

Rats were dark-adapted overnight and prepared for recording under red illumination (LED, >620 nm). The individual drug experiments were performed on 11 rats of similar ages. They were initially anesthetized with an intramuscular injection of 86 mg/kg ketamine and 6.5 mg/kg xylazine into the hamstring muscles. For recording, animals were placed on a thermostatically controlled electrical heating blanket (CWE, Inc., Ardmore, PA) and anesthesia was maintained with half the initial dose every 45-60 min via a subcutaneous needle fixed in the flank. Rectal temperature was maintained at 36-37 °C. Animals generally recovered after the sessions; however, after they were given intravitreal injections in both eyes, they were killed using 100 mg/kg, intraperitoneal injection of sodium pentobarbital. Pupils were dilated to 5 mm in diameter with topical 0.5% atropine and 2.5% phenylephrine. A drop of proparacaine hydrochloride (0.5%) was used for corneal anesthesia. The animal's head was held steady, in order to reduce noise originating from respiratory and other movements, using an aluminum head holder with a hole for the upper incisors. This head holder also served as the ground.

Recording sessions lasted 4 to 8 h. ERGs were recorded differentially between Dawson, Trick, and Litzkow (DTL) fiber electrodes [43], moistened with saline and placed on both eyes. The stimulated eye was covered with a clear contact lens heat molded from ACLAR (Ted Pella, Inc., Redding, CA), and the control eye was covered with an opaque plastic contact lens. Both lenses were placed over a cover of 1% methylcellulose sodium (Celluvisc, Allergan Inc., Irvine, CA). Signals were amplified (DC to 300 Hz), digitized at 1 KHz with a resolution of 2 μV, and sent to the computer for averaging, display, storage, and subsequent analysis.

ERGs were recorded using brief full field flashes (λmax, 462 nm; -5.8 to 1.9 log sc td s) under fully dark-adapted conditions, in most cases, following a conditioning flash, to suppress rod-driven activity as described in Results. Intervals between flashes were adjusted so that the response returned to baseline before another stimulus was presented. A digital 60 Hz notch filter was applied offline.

Stimulus calibration was performed by measuring scotopic luminance with a scotopically corrected (for humans) photometer (International Light, model IL1700, Newburyport, MA) [44]. The LEDs in the stimulator were activated with a 1 kHz train of current pulses each having a duration of 100 μs and the luminance was measured. The luminance-time product for each of the single flash stimuli was expressed in sc cd s m-2 and converted to sc td s by multiplying the product by the pupil area.

Intravitreal injections

Intravitreal injections were performed under a binocular operating microscope (magnification: 10x), under dim red illumination (>620 nm). A small hole (about 0.5 to 1 mm) was punctured behind the limbus using a 27 gauge needle to allow penetration at 45 °C of the underlying sclera, choroid, and retina by a fine glass micropipette (tip about 20 μm; diameter: 1 mm) fixed on a 10 μl Hamilton microsyringe (Hamilton Company, Reno, NV) for injecting pharmacological agents. A single about 2 μl injection was given slowly over 1 min in the vitreous. The intravitreal drug concentrations used were about 3 μM TTX and 200 μM 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX), based on an estimated vitreous chamber volume of 40 μl in the rat [45,46]. After injection, the ERG was monitored until no further change in response was seen, generally about one hour, before taking data.

Electroretinogram analysis

ERG amplitudes before and after drug administration were measured at a fixed time of 50 ms after the stimulus flash, corresponding to the time of peak of the cone-driven b-wave and plotted against the flash energy to generate stimulus response curves.

Results

Nav1 α-subunit epitope sequence analysis and specificity

The anti-Nav1 α-subunit-specific monoclonal antibodies used in this study were generated using rat-specific peptides located within C-terminus (Nav1.1 and Nav1.2) or a non-conserved region near the sixth transmembrane domain (Nav1.6; Figure 1A). Because we were also using a more distantly related species (rabbit) that was not used in previous studies, we wanted to predict whether these monoclonal antibodies would recognize orthologous Nav1 α-subunit proteins. To this end, we used a bioinformatics approach to examine sequence conservation of these peptide immunogens in Nav1.1, Nav1.2 and Nav1.6 across mammalian species (Table 3). To identify the most highly conserved sequences, we used AlignX software to do pair-wise alignments of each of the peptides with the orthologous Nav1 α-subunit proteins from each species and calculated the percent amino acid identity. We found that the Nav1.1, Nav1.2, and Nav1.6 peptide immunogens were each highly conserved among rat, mouse, rabbit, dog, bovine, and human proteins (Figure 2). Nav1.1 and Nav1.2 peptides were 94-100% identical with their respective orthologs across all species and the shorter Nav1.6 peptide was 100% identical between rat and mouse and 88-94% between rat and non-rodent Nav1.6 sequences. Based on this high level of sequence identity, each of the monoclonal antibodies was predicted to recognize their orthologous proteins in mammals.

The first 36 amino acids of the Nav1.1 and Nav1.2 peptides differed at only three residues (Figure 2B); therefore, we did pair-wise alignments of each peptide immunogen with all members of the Nav1 family in rat to determine the extent of this sequence similarity (Table 1). These analyses revealed that over its entire length, the Nav1.2 peptide immunogen, respectively shared 64%, 73%, and 71% amino acid identity with rat Nav1.1, Nav1.3, and Nav1.7. Although this level of sequence conservation raises the possibility that the anti-Nav1.2 monoclonal antibody could potentially crossreact with these other Nav1 family proteins, this is unlikely. If the anti-Nav1.2 antibody recognizes a portion of the epitope that is highly conserved across these Nav1 family members, then the Nav1.2 antibody would label all structures or cells that express Nav1.1, Nav1.3, or Nav1.7. However, this is not the case, as the Nav1.2 antibody produces immunolabeling patterns distinct from other Nav1 family members. The Nav1.2 immunolabeling pattern is distinct from the Nav1.1 pattern in rat cerebellar granule cells [47] and axon initial segments of Purkinje neurons in Nav1.6 null mice [15] and differential labeling by anti-Nav1.2 and Nav1.3 antibodies has been reported in the myenteric nervous system [48]. Therefore, despite the modest sequence conservation of the amino acid sequence in the peptide used to raise the Nav1.2 antibody, the antibody is unlikely to cross react with other Nav1 α-subunit family members. Direct testing of the Nav1.2 antibody shows no crossreactivity with Nav1.1, Nav1.3, or Nav1.6 (Dr. James Trimmer, personal communication).

As an additional evaluation of the specificity of the anti-Nav1 α-subunit antibodies, we used peptide immunogen sequences for BlastP and tBlastn searches of the SwissProt, non-redundant and EST databases (NCBI) to determine if they shared sequence identity with other known or predicted mammalian proteins. In Blast searches, each match is scored by an E-value that indicates the number of matches predicted to occur by chance alone, therefore, we examined all matches with an E value of less than 1 [42]. For each of the searches, all matches identified were for Nav1 family proteins. Therefore, no other mammalian proteins were predicted to contain the peptide epitopes used to generate these monoclonal antibodies, consistent with our conclusions that they were specific for the different Nav1 α-subunit proteins.

Localization of Nav1 α-subunit

We first assessed the overall distribution of Nav1 α-subunits in the retina in comparison to the distribution of Nav channels reported in previous physiological and anatomical studies. These studies have clearly established the presence of Nav1 α-subunits in third-order amacrine and ganglion cells, as discussed in the Introduction. Each Nav1 α-subunit antibody used in these studies produced the expected labeling of amacrine and ganglion cells. In addition, each Nav1 α-subunit antibody produced a characteristic pattern of labeling that included cells and processes in the distal retina, extending the findings of the previous studies.

Pan-Nav labeling

Labeling using the Pan-Nav antibody revealed a characteristic pattern of Nav1 α-subunit distribution that was similar in rat and mouse retina (Figure 3, rat retina shown). In vertical cryosections of retina, Pan-Nav labeling was present in ganglion cell bodies and axons and in processes in the inner plexiform layer (IPL), consistent with previous studies (Figure 3A,B). Two distinct sets of cell bodies in the inner nuclear layer (INL) were also labeled. One set of cells corresponded to amacrine cells located near the border of the IPL and INL. The second set consisted of large flat cells at the distal edge of the INL that corresponded to horizontal cells (see next section). Little Pan-Nav immunoreactivity was observed in the mid-INL, where the cell bodies of bipolar and Müller glial cells reside, or in the outer nuclear layer (ONL), where the photoreceptors reside. Relatively little Pan-Nav labeling was present in the IPL or outer plexiform layer (OPL) in frozen sections. However, discretely labeled processes similar to the "morphologically distinct processes" described previously [14] were observed.

Labeling using the Pan-Nav antibody was highly sensitive to fixation and processing of the tissue and this was particularly labile in the plexiform layers. For frozen sections of retina that were fixed for more than 15 min total, antigen retrieval procedures were required to obtain Pan-Nav labeling. In contrast, wholemounted retinas that were only lightly fixed (5 min) and never subjected to freezing did not require antigen retrieval for immunostaining with the Pan-Nav antibody (Figure 3C-H, single optical planes focused at different retinal levels shown). Immunolabeling of lightly fixed rat retinal wholemounts confirmed the pattern of Nav1 α-subunit distribution observed in frozen sections and revealed additional, fixation-sensitive immunoreactivity that was not detected in frozen sections. Pan-Nav labeling was found in the cell bodies of ganglion, amacrine, and horizontal cells, similar to labeling seen in vertical frozen sections. The identity of these cell types was confirmed by double labeling for Pan-Nav and the markers calbindin and calretinin which label ganglion, amacrine and horizontal cells (data not shown). Ganglion cell axons and axon initial segments showed strong Pan-Nav labeling, as expected. Much more pronounced Pan-Nav labeling was present in the plexiform layers in wholemounts than in frozen sections. In the OPL of retinal wholemounts, small puncta of Pan-Nav labeling were embedded on a background of more diffuse labeling. The IPL of retinal wholemounts showed abundant diffuse Pan-Nav labeling at all levels in addition to intensely labeled varicose processes similar to the morphologically distinct processes described previously. Photoreceptor cell bodies in the ONL and bipolar and Müller cell bodies in the mid-INL showed scant Pan-Nav labeling, similar to results seen in cryosections.

Labeling with isoform-specific Nav1 α-subunit antibodies

Each isoform-specific Nav1 α-subunit antibody produced a characteristic labeling pattern consistent with the overall distribution of Nav1 α-subunit shown by the Pan-Nav antibody (Figure 4; mouse retina illustrated). In contrast to Pan-Nav antibody, labeling using the isoform-specific Nav1 α-subunit antibodies was not fixation-sensitive and did not require antigen retrieval.

Nav1.1

As expected, labeling for Nav1.1 was found in cell bodies in the GCL, ganglion cell axons, a subpopulation of amacrine cells along the INL/IPL border and strongly labeled processes in the IPL (Figure 4A). In addition, Nav1.1 immunoreactivity was found in the OPL, although little labeling was present in bipolar or Müller cell bodies in the mid-INL or photoreceptor cell bodies in the ONL. A similar distribution of Nav1.1 labeling was seen in wholemounts, which also revealed strong labeling of structures similar in appearance to ganglion cell axon initial segments (Figure 4D).

Nav1.2

Labeling for Nav1.2 also was distributed as expected in the inner retina (Figure 4B). Cell bodies in the GCL and along the INL/IPL border showed labeling. Although some strongly labeled processes were observed in the IPL, labeling tended to be more diffuse, and strongly labeled processes were seen less often than with Nav1.1 labeling. In addition to labeling in the inner portion of the retina, strong Nav1.2 labeling was found in a set of cell bodies located along the OPL/INL border and diffuse labeling was present in the OPL. Little Nav1.2 labeling was found in the mid-INL or ONL. Labeling for Nav1.2 in wholemounts was similar, although labeling of ganglion cell axons and of structures with the appearance of ganglion cell axon initial segments was more obvious than in vertical sections (Figure 4E).

Nav1.6

The pattern of Nav1.6 labeling also was consistent with the labeling pattern seen with the Pan-Nav antibody (Figure 4C). As expected, Nav1.6 immunolabeling was found in cell bodies in the GCL. Nav1.6 labeling was also seen in a subset of amacrine cells at the INL/IPL border and diffusely distributed in the IPL. Ganglion cell axons also showed Nav1.6 labeling as appropriate, particularly structures with the appearance of ganglion cell axon initial segments (Figure 4F). Labeling for Nav1.6 also was present in the OPL and a population of horizontal cells along the distal border of the INL. There was little Nav1.6 labeling in the mid-INL or ONL, similar to the Pan-Nav, Nav1.1, and Nav1.2 labeling patterns.

Together, these results confirm that amacrine and ganglion cells, their processes in the IPL and ganglion cell axons, express Nav1.1, Nav1.2 and Nav1.6 α-subunit, consistent with Pan-Nav labeling and previous reports [13-15]. These results also indicated that there was little expression of Nav1 α-subunits in the cell bodies of photoreceptors in the ONL or bipolar and Müller cells in the mid-INL, although labeling was present in the OPL and a population of cells along the OPL/INL border.

Nav1 α-subunit immunolabeling was not detected in bipolar cell bodies

Recent patch-clamp studies have indicated the presence of functional Nav channels in isolated rat bipolar cells [24,49], although we observed little labeling for any Nav1 α-subunit in the mid-INL where bipolar cell bodies reside. To better understand the distribution of Nav1 α-subunits and their potential functional roles in bipolar cells, we performed triple labeling in the rat retina using the Pan-Nav antibody in conjunction with antibodies against protein kinase C and Chx-10 that label rod bipolar cells or all types of bipolar cells, respectively [50,51]. These studies definitively showed that the cell bodies along the distal border of the INL showing Nav1 α-subunit labeling were not bipolar cells and that the Nav1 α-subunit-positive cells in the innermost portion of the INL represented amacrine cell populations. No colocalization of Pan-Nav labeling with either protein kinase C or Chx10 labeling was present, indicating that Nav1 α-subunits were either absent from bipolar cell bodies or expressed at a level too low to detect (Figure 5).

Horizontal cells and other cell types ramifying in the outer plexiform layer express Nav1 α-subunits

The finding that the cells in the distal INL showing Nav1 α-subunit labeling did not express bipolar cell markers suggested that these cells might be horizontal cells. To test this, we performed double labeling for Pan-Nav and calbindin, a marker for horizontal cells in rat, mouse and rabbit retina [52-54]. Horizontal cell bodies and processes showed Pan-Nav labeling in rat, mouse, and rabbit retina (Figure 6). However, additional diffuse Pan-Nav labeling also was present throughout the OPL that did not correspond to horizontal cell processes, indicating that Nav1 α-subunit expression in the OPL was not limited strictly to horizontal cells. In the rabbit OPL specifically, discrete annular clusters of Pan-Nav labeling were present in addition to diffuse labeling. Together, these results indicate that Nav1 α-subunits were present on horizontal cells in addition to the processes of other cell types that ramify in the OPL.

Nav1 α-subunit isoforms are differentially distributed within horizontal cells and the outer plexiform layer

To better understand the distribution of Nav1 α-subunits in the outer retina, we performed a series of double labeling experiments to characterize further the distribution of Nav1.1, Nav1.2, and Nav1.6 subunits within horizontal cells and the OPL.

In both rat and mouse retinas, horizontal cell bodies and processes showed scant Nav1.1 labeling, while appreciable Nav1.1 labeling was present in the OPL distal to the horizontal cell processes (Figure 7A-C; rat shown). Rabbit horizontal cell bodies and processes, however, showed distinct labeling for Nav1.1 (Figure 7D-F). Labeling for Nav1.1 in the rabbit OPL also showed discrete annular clusters similar to those observed in the rabbit OPL using the Pan-Nav antibody. The Nav1.1 clusters in the rabbit OPL did not colocalize with calbindin, suggesting that they arose from cells other than horizontal cells, consistent with the pattern seen using the Pan-Nav antibody.

Nav1.2 and Nav1.6 labeling was closely associated with horizontal cells in all three species, colocalizing extensively with calbindin labeling and with comparatively little additional labeling for Nav1.2 or Nav1.6 detected elsewhere in the OPL. Horizontal cell bodies and processes, including the dendritic tips, showed prominent Nav1.2 labeling (Figure 7G-I; rat shown). In contrast, labeling for Nav1.6 in horizontal cells was appreciably stronger in the cell body and proximal processes than in the dendritic tips (Figure 7J-L; rat shown). Nav1.2 and Nav1.6 labeling patterns in the horizontal cells and OPL were similar in the retinas of rat, mouse, and rabbit.

These results indicate that Nav1.1, Nav1.2, and Nav1.6 α-subunits are all expressed by mammalian horizontal cells. Although horizontal cells expressed Nav1.1, much of the Nav1.1 labeling in the OPL did not colocalize with calbindin, suggesting that Nav1.1 also was present on processes from other cell types. In contrast, Nav1.2 and Nav1.6 in the distal retina were closely associated with horizontal cells and their processes, indicating that other cell types ramifying in the OPL expressed little Nav1.2 and Nav1.6.

Nav1.1 is diffusely expressed by multiple cell types in the outer plexiform layer

Imaging of rat retinal wholemounts double-labeled for Nav1.1 and calbindin confirmed that much of the diffuse Nav1.1 labeling in the OPL was not localized to horizontal cell processes themselves (Figure 8A-C; single optical section shown). The presence of additional Nav1.1 labeling in the OPL that did not correspond to horizontal cell processes indicated that Nav1.1 was expressed by other cell types ramifying in the OPL, such as bipolar cells or photoreceptors. To better understand the distribution of Nav1.1 in the OPL, we performed double labeling for Nav1.1 and markers for photoreceptor terminals and the tips of ON bipolar cell dendrites and examined them by confocal microscopy in single optical sections.

To determine whether Nav1.1 labeling might be associated with photoreceptor terminals, we double labeled sections for Nav1.1 and vesicular glutamate transporter 1 (VGluT1), which is expressed in both rod and cone terminals [39,55]. Examination of single optical sections in the OPL showed appreciable overlap of Nav1.1 and VGluT1 labeling, suggesting that some Nav1.1 labeling in the OPL was associated with photoreceptor terminals (Figure 8D-F). However, much of the Nav1.1 labeling in the OPL did not colocalize with photoreceptor terminals.

The presence of Nav1.1 labeling that did not colocalize with either horizontal cell or photoreceptor terminals suggested that Nav1.1 also might be expressed on the processes of other cells ramifying in the OPL. To test whether Nav1.1 was associated specifically with ON-cone bipolar and rod bipolar cell dendrites at the point of contact with the synaptic ribbons of cone and rod terminals, we double labeled sections for Nav1.1 and mGluR6, the glutamate receptor on ON-cone and rod bipolar cell dendrites at ribbon synapses [56], and examined them by confocal microscopy. Examination of single optical planes (approximately 0.3 μm thickness) revealed that Nav1.1 and mGluR6 labeling showed little colocalization, although they were often found in close proximity (Figure 8G-I). Thus, Nav1.1 did not specifically colocalize with mGluR6 on bipolar cell dendrites within the ribbon synapse complexes.

Effects of tetrodotoxin on the rat electroretinogram

Given the extensive labeling for Nav α-subunits in horizontal cells and the OPL, we were interested in looking more closely at the effects on outer retinal function of blocking retinal Nav channels. Similar to past studies in rats [57], intravitreal injections of TTX to block Nav channels severely attenuated the potentials in the dark-adapted ERG known to arise from amacrine cells and retinal ganglion cells (p-STR and n-STR; Figure 9B,). Past studies indicate that TTX does not alter the amplitude of waves of the dark-adapted ERG originating more distally, the a-wave from rods and the b-wave from rod bipolar cells. However, under photopic conditions, TTX reduces the amplitude of the cone-driven b-wave, a signal generated primarily by cone bipolar cells [57]. In the present study we recorded ERGs from rats under dark-adapted conditions, but with rod signals active or suppressed to study more closely the effects of TTX on the rod vs. cone circuits under similar conditions of light-adaptation (Figure 9).

ERG responses to low stimulus energies up to those that saturated the dark-adapted b-wave (e.g., response to -1.3 sc td s in Figure 9A), are believed to originate from the sensitive rod to rod bipolar cell circuit (e.g., [58-60]). Following TTX blockade, we observed, that the b-wave was unaffected over the range of these stimulus intensities, as illustrated here for a representative result (unpublished observations). This finding indicates that cells of origin of the b-wave in the rod to rod bipolar cell circuit were hardly affected by TTX. B-waves in response to higher stimulus energies in the dark-adapted ERG (1.3 and 1.6 log sc td; Figure 9A) were reduced by TTX. Dark-adapted ERG responses to strong stimuli were mixed rod-cone responses (e.g., [58,61]), meaning cone circuits were active.

We investigated the hypothesis that the TTX-mediated attenuation of the b-wave was occurring in cone circuits in the mixed rod-cone ERG by isolating dark-adapted cone-driven responses using a cone isolation protocol. Briefly, as described in studies by other investigators [62,63], an initial conditioning flash (1 s duration) was used to suppress the rod photocurrent and then, at an appropriate time interval for recovery of cone but not rod responses, a test flash was presented [61,62,64,65]. In rodents, an 800 ms interval between conditioning and test flashes allows full recovery of cone signals while rod signals are still suppressed [62,66]. Using an interstimulus interval of 800 ms, we were able to isolate and study dark-adapted cone-driven responses over a range of stimulus energies. As shown in Figure 9C, the isolated cone-driven b-wave was reduced by TTX for all stimulus energies that elicited a response (more fully documented in unpublished observations). The effect on the cone-driven response was more than sufficient to account for the reduction in the dark-adapted mixed rod-cone ERG.

In the next phase of the experiment, the dark-adapted cone-isolation protocol was repeated following blockade with CNQX of α-amino-5-hydroxy-3-methyl-4-isoxazole propionic acid (AMPA) and kainate (KA) receptors [67]. In the retina, these receptors are located on the dendrites of horizontal, OFF bipolar, amacrine, and ganglion cells. Finally, the experiment was repeated again after intravitreal injection of TTX in the eyes already injected with CNQX (Figure 10). Representative ERG traces from a single animal (Figure 10A) show that CNQX did not affect the cone-driven ERG. In contrast, CNQX+TTX produced a large reduction of b-wave amplitude. B-wave amplitudes were measured at a fixed time, (50 ms; the cone-driven b-wave time to peak), and plotted as a function of flash energy (Figure 10B). The plot confirms that blockade of AMPA/KA receptors with CNQX left the average b-wave amplitude unchanged, whereas CNQX+TTX produced a large attenuation of the b-wave (n= 6 eyes from 4 rats). The average percent reduction of the maximum b-wave amplitude after CNQX+TTX was very similar to that produced by TTX alone in other eyes (CNQX+TTX: 59%±14.3%, n=6, this study; TTX: 67.6%±17.3%, n=7 eyes in 7 rats, unpublished observations). These results indicate that the blockade of AMPA/KA receptors on neurons in cone circuits located proximal to photoreceptors did not alter the TTX-mediated attenuation of the cone-driven b-wave and suggest that the effect of the TTX was directly on Nav channels in cone bipolar cells. The nearly parallel downward shift of the b-wave energy-amplitude function in Figure 10B after TTX was injected indicates an effect on response amplitude, more than on sensitivity. This also is consistent with an effect on Nav channels in cone bipolar cells, rather than the cone photoreceptors that provide input to the bipolar cells. Some of these results have been published previously in abstract form [66].

Discussion

Previous studies in the rat retina have shown specific distributions of Nav1.1, Nav1.2 and Nav1.6 in the retinal ganglion cells and their axons [12,13,15] and in the IPL [14]. Our studies confirm these findings in the inner retina and demonstrate that each of these Nav1 α-subunits also are present more distally in the retina, particularly in the OPL and horizontal cells. In addition, while Nav1.1, Nav1.2 and Nav1.6 show substantial overlap in our study, the distribution of each subunit in the retina is unique.

A striking finding in our studies is the localization of Nav1 α-subunits to the OPL and horizontal cells and their processes. The localization of Pan-Nav labeling and labeling for Nav1.1, Nav1.2 and Nav1.6 α-subunits to rat, mouse, and rabbit horizontal cells is consistent with a previous report of Nav1 α-subunit immunolabeling in horizontal cells in the cat and monkey retina [68]. Furthermore, the current findings are consistent with previous functional studies demonstrating TTX-sensitive Na+ currents in isolated horizontal cells in several species: catfish: [30]; white perch: [31], cat: [32], rabbit: [33].

Nav1 α-subunit labeling of horizontal cells was most prominent in the cell body, but was also present in the processes. Our studies using antibodies directed against specific Nav1 α-subunits indicate that rat, mouse, and rabbit horizontal cells express at least two types of Nav1 α-subunits: Nav1.2 and Nav1.6. Additionally, rabbit horizontal cells show substantial labeling for Nav1.1. Co-expression of multiple Nav1 α-subunit within individual neurons has been reported previously in retinal ganglion cells [13,15] and neurons in the adult brain [69-71]. All horizontal cells in the rat and mouse retina showed similar expression of Nav1.2 and 1.6, consistent with the presence of only a single type of axon-bearing horizontal cell in the rodent retina [54]. In contrast, rabbit horizontal cells did not show uniform labeling for Nav1 α-subunits, suggesting heterogeneous Nav1 α-subunit expression among rabbit horizontal cells (Figure 6J-L and Figure 7D-F). The large size of the Nav1 a immuno-positive horizontal cells in rabbit suggests that at least the axonless A-type horizontal cells that exclusively contact cones in the rabbit retina express Nav1 α-subunits [72,73].

The strong labeling of the processes and cell bodies of horizontal and amacrine cells is consistent with somatic labeling of neurons in the adult mammalian brain reported previously [74,75]. Voltage-gated Na+ channels containing Nav1.1 in the somato-dendritic compartment of neurons in rat brain have been proposed to play a key role dendritic excitability [75]. Somato-dendritic labeling also may represent cytoplasmic Nav1 α-subunit intermediates in the biosynthetic pathway [76] or storage pools. The intensity of Pan-Nav labeling seen in horizontal and amacrine cell bodies also may reflect the expression of more than one Nav1 α-subunit in these cells. In addition, the cells may possess other Nav1 α-subunits that we did not test.

The results of our single and double immunolabeling analyses suggest that Nav1 α-subunits in the OPL are not restricted only to horizontal cells. Pan-Nav labeling was diffusely distributed throughout the OPL, consistent with the band of Nav1.1 labeling found mainly distal to the large processes of the horizontal cells and the weaker, diffuse labeling for Nav1.2 and Nav1.6 present in the OPL. This diffuse Nav α-subunit labeling could not be accounted for by the processes of any single cell type ramifying in the OPL, suggesting that Nav α-subunits may be present on the processes of multiple cell types that ramify in the OPL.

Our data suggest that some of the additional Nav α-subunit labeling in the OPL is associated with photoreceptor terminals. Double labeling for Nav1.1 and VGluT1, a photoreceptor terminal marker [39,55], indicates that at least Nav1.1 is present at the level of the photoreceptor terminals. The patchy distribution of Nav1.1 labeling in the OPL of flatmounted retinas would be consistent with the spatial arrangement of cone terminals (see Figure 8A-C). However, the limits of optical resolution in our confocal images (approximately 0.3 μm), the anatomical complexity of the processes in and around the photoreceptor terminals, and the diffuse nature of the labeling preclude unequivocal assignment of Nav1.1 labeling at the level of the photoreceptor terminals to specific cell types. Other studies also support expression of functional voltage-gated Na+ channels by photoreceptors. Patch clamp recordings of human photoreceptors in isolation or in retinal slices have identified TTX-sensitive Na+ currents [35,36] and knockout of the gene for Nav1.6 in mouse has been shown to impede the physiological maturation of photoreceptors [6]. Interestingly, in human photoreceptors, single-cell PCR showed Nav1.2-rather than Nav1.1-transcripts [28,36].

Although Nav α-subunits were clearly present in horizontal cells and at the level of photoreceptor terminals, these structures could not account for all Nav α-subunit labeling in the OPL. One distinct possibility is that some of the diffuse Nav α-subunit labeling was associated with bipolar cell dendrites. Several pieces of evidence support this possibility: depression of the cone-driven b-wave of the ERG, which arises primarily from bipolar cells, by TTX blockade of Nav channels (this study); patch-clamp recordings of voltage-gated sodium currents in individual bipolar cells [24,25,28,49], and previous reports of Nav channel gene or protein expression by bipolar cells [25,28,68]. The diffuse nature of the labeling and the limits of detection and anatomical resolution associated with the immunolabeling techniques in the current study do not permit the direct assignment of Nav α-subunits to bipolar cell dendrites. However, the current studies do suggest that Nav α-subunits are not concentrated at the tips of bipolar cell dendrites contacting photoreceptor ribbons as there was little colocalization of Nav α-subunit and mGluR6 labeling. The current studies also indicate that Nav α-subunits are not present in large quantities in the cell bodies of bipolar cells.

An interesting finding of our study was that the extent of labeling by the Pan-Nav antibody was highly sensitive to fixation time and processing of the tissue, especially in the plexiform layers. This indicates that increased fixation and processing interferes with immunodetection of Nav1 α-subunits, when the Pan-Nav antibody is used. The Nav1 α-subunit-specific antibodies did not show the same sensitivity to fixation, although labeling in the lightly fixed wholemounts was stronger than in frozen sections. Antigen retrieval procedures were required to obtain Pan-Nav labeling in frozen sections that had been exposed to fixative for as short as 30 min. Even with antigen retrieval procedures, Pan-Nav labeling in the plexiform layers in frozen sections remained limited. In contrast, in wholemounts that were lightly fixed for only five minutes and were never subjected to freezing, Pan-Nav labeling was intense and readily detectible in the plexiform layers and neuronal cell bodies without antigen retrieval. The α-subunit-specific antibodies also produced high levels of labeling in the plexiform layers, consistent with the high levels of Pan-Nav labeling seen in the lightly fixed wholemounts. Together, these results suggest that the distribution of Nav1 α-subunits in the plexiform layers is much wider than has been appreciated previously and that use of light fixation enhances immunohistochemical detection of Nav1 α-subunits.

It is unlikely that labeling for Nav1 α-subunits in the outer retina, neuronal cell bodies and the plexiform layers is due to non-specific labeling. The specificity of the antibodies used in the current study has been extensively characterized using immunoblotting, ELISA and preadsorption assays, and immunolabeling of cells and tissues known to either contain or lack specific Nav1 α-subunits [13,14,37]; this report. Our further analysis of epitope sequence homology across Nav1 α-subunits and other mammalian protein sequences showed that these epitopes are unique to Nav1 α-subunits and, in the case of the α-subunit specific antibodies, are isoform specific as well. The Pan-Nav, Nav1.1, Nav1.2, and Nav1.6 labeling patterns observed in our studies precisely corroborate previous reports localizing these subunits in the inner retina [12-15]. Finally, Nav1 α-subunit labeling patterns in the outer retina are consistent with emerging physiological and molecular data indicating that Nav1 α-subunits and functional Nav channels are present in outer retina [24-26,28,35,36,49].

The dark-adapted rod-driven a-waves were unaffected by TTX, implying that rod photocurrents were unaffected (Figure 9 and unpublished observations). Cone photoreceptor contributions to a-waves were negligible in our experiments, making it difficult to evaluate effects of TTX on cone photocurrents, but it is unlikely that the photocurrents were affected. Labeling for Nav1 α-subunit has not been observed in photoreceptor outer segments where photoreceptor a-waves are generated. However, the dark-adapted cone-driven b-waves were reduced in amplitude by TTX, raising the possibility that there were effects on cone terminals, cone bipolar cells, or other components of the cone circuits. In addition to the ON cone bipolar cells, the cone-driven b-wave also can be shaped by the activity of OFF cone bipolar cells, horizontal cells, as well as more proximal retinal neurons and Müller cells [77-79]. Blockade of inputs to these additional neuronal contributors to the b-wave by CNQX did not alter the cone-isolated ERGs. This reduces the likelihood that TTX effects were mediated by horizontal cells or any of the other neurons in the cone circuit with synaptic input via AMPA/KA receptors. However, the b-wave amplitude after CNQX+TTX was reduced and the amount of attenuation was similar to that caused by TTX alone. From these data, we infer that the ON cone bipolar cells themselves are the most likely locus of the TTX effects. Cone inner segments or terminals are also a logical possibility. TTX-sensitive currents have been observed in isolated human photoreceptors, but predominantly in rods (68% of those studied) rather than cones (19%) [35,36]. However, the physiological role of these currents in light-evoked signaling is unclear and in response to flashed stimuli, photoreceptor membranes would be too hyperpolarized to allow Nav channels to open [80], making it unlikely that cones contribute to the effects of TTX reported here. The ERG results would be consistent with the presence of Nav channels in the dendritic compartment of ON cone bipolar cells. Past studies also indicate that TTX-sensitive Na+ currents are present in the somato-dendritic compartment of the rat cone bipolar cells [24].

The role of Nav channels in horizontal cells is puzzling. Under dark-adapted conditions horizontal cells undergo tonic depolarization by glutamate release from the photoreceptors that keep the resting membrane potential around -35 mV (for review see [81]), thereby causing voltage-dependent inactivation of any Nav channels. A large hyperpolarization of the horizontal cells due to a high energy flash might potentially relieve this inactivation. Voltage-gated Na+ channels could allow inward Na+ currents in horizontal cells after offset of a strong stimulus when the membrane repolarizes [82,83]. The impact of this in retinal processing would need further study.

The role of Nav channels in mammalian ON-cone bipolar cells also needs elucidation. It has been suggested that Nav channels in bipolar cells may augment subthreshold depolarizing potentials in the dendrites and cell body of bipolar cells [25] in a manner similar to those seen in CNS neurons [84]. The Nav channels may play a special role in circuits carrying transient signals through the retina [26,27]. Our results in other studies suggest that the TTX-sensitive augmentation of the cone-driven b-wave is maximal under the dark-adapted conditions used in the present study and decreases with increasing background illumination (unpublished observations). This result is similar to the finding in salamander retina, where TTX-sensitive Na+ currents in transient ON bipolar cells were maximal under the dark-adapted condition and decreased with light adaptation [85]. Future studies will provide more insight about the diverse roles that Nav channels play in retinal processing.

Acknowledgements

We thank Drs. Gary Matthews, James Trimmer, and Costa Colbert for their generous gifts of antibodies and helpful comments. Monoclonal antibodies K74/71, K69/3, and K87A/10 were developed by as well as obtained from the University of California Davis/NINDS/NIMH NeuroMab Facility, supported by NIH grant U24NS050606 and maintained by the Department of Pharmacology, School of Medicine, University of California, Davis, CA 95616. This study was supported by NIH grants EY06671 (L.J.F.) and EY07551 (U.H.C.O.).

References

1. Yu FH, Catterall WA. Overview of the voltage-gated sodium channel family. Genome Biol 2003; 4:207.

2. Yu FH, Catterall WA. The VGL-chanome: a protein superfamily specialized for electrical signaling and ionic homeostasis. Sci STKE 2004; 2004:re15.

3. Hodgkin AL, Huxley AF. Propagation of electrical signals along giant nerve fibers. Proc R Soc Lond B Biol Sci 1952; 140:177-83.

4. Makielski JC, Farley AL. Na(+) current in human ventricle: implications for sodium loading and homeostasis. J Cardiovasc Electrophysiol 2006; 17 Suppl 1:S15-S20.

5. Catterall WA. From ionic currents to molecular mechanisms: the structure and function of voltage-gated sodium channels. Neuron 2000; 26:13-25.

6. Cote PD, De Repentigny Y, Coupland SG, Schwab Y, Roux MJ, Levinson SR, Kothary R. Physiological maturation of photoreceptors depends on the voltage-gated sodium channel Nav1.6 (Scn8a). J Neurosci 2005; 25:5046-50.

7. Isom LL. Sodium channel beta subunits: anything but auxiliary. Neuroscientist 2001; 7:42-54.

8. Goldin AL. Diversity of mammalian voltage-gated sodium channels. Ann N Y Acad Sci 1999; 868:38-50.

9. Goldin AL. Resurgence of sodium channel research. Annu Rev Physiol 2001; 63:871-94.

10. Wollner DA, Catterall WA. Localization of sodium channels in axon hillocks and initial segments of retinal ganglion cells. Proc Natl Acad Sci U S A 1986; 83:8424-8.

11. Fjell J, Dib-Hajj S, Fried K, Black JA, Waxman SG. Differential expression of sodium channel genes in retinal ganglion cells. Brain Res Mol Brain Res 1997; 50:197-204.

12. Boiko T, Rasband MN, Levinson SR, Caldwell JH, Mandel G, Trimmer JS, Matthews G. Compact myelin dictates the differential targeting of two sodium channel isoforms in the same axon. Neuron 2001; 30:91-104.

13. Boiko T, Van Wart A, Caldwell JH, Levinson SR, Trimmer JS, Matthews G. Functional specialization of the axon initial segment by isoform-specific sodium channel targeting. J Neurosci 2003; 23:2306-13.

14. Van Wart A, Boiko T, Trimmer JS, Matthews G. Novel clustering of sodium channel Na(v)1.1 with ankyrin-G and neurofascin at discrete sites in the inner plexiform layer of the retina. Mol Cell Neurosci 2005; 28:661-73.

15. Van Wart A, Matthews G. Impaired firing and cell-specific compensation in neurons lacking nav1.6 sodium channels. J Neurosci 2006; 26:7172-80.

16. Lipton SA, Tauck DL. Voltage-dependent conductances of solitary ganglion cells dissociated from the rat retina. J Physiol 1987; 385:361-91.

17. Skaliora I, Scobey RP, Chalupa LM. Prenatal development of excitability in cat retinal ganglion cells: action potentials and sodium currents. J Neurosci 1993; 13:313-23.

18. Schmid S, Guenther E. Developmental regulation of voltage-activated Na+ and Ca2+ currents in rat retinal ganglion cells. Neuroreport 1996; 7:677-81.

19. Boos R, Schneider H, Wassle H. Voltage- and transmitter-gated currents of all-amacrine cells in a slice preparation of the rat retina. J Neurosci 1993; 13:2874-88.

20. Smith RG, Vardi N. Simulation of the AII amacrine cell of mammalian retina: functional consequences of electrical coupling and regenerative membrane properties. Vis Neurosci 1995 Sep-Oct; 12:851-60.

21. Feigenspan A, Gustincich S, Bean BP, Raviola E. Spontaneous activity of solitary dopaminergic cells of the retina. J Neurosci 1998; 18:6776-89.

22. Hartveit E. Reciprocal synaptic interactions between rod bipolar cells and amacrine cells in the rat retina. J Neurophysiol 1999; 81:2923-36.

23. Cohen ED. Voltage-gated calcium and sodium currents of starburst amacrine cells in the rabbit retina. Vis Neurosci 2001 Sep-Oct; 18:799-809.

24. Pan ZH, Hu HJ. Voltage-dependent Na(+) currents in mammalian retinal cone bipolar cells. J Neurophysiol 2000; 84:2564-71.

25. Zenisek D, Henry D, Studholme K, Yazulla S, Matthews G. Voltage-dependent sodium channels are expressed in nonspiking retinal bipolar neurons. J Neurosci 2001; 21:4543-50.

26. Ichinose T, Shields CR, Lukasiewicz PD. Sodium channels in transient retinal bipolar cells enhance visual responses in ganglion cells. J Neurosci 2005; 25:1856-65.

27. Saszik SM, DeVries SH. A TTX-Sensitive Na+-Current in One Type of On Bipolar Cell in the Ground Squirrel Retina. Invest Ophthalmol Vis Sci 2005; 46: ARVO E-Abstr. 1122.

28. Miyachi E-I, Kawai F, Ohkuma M, Horiguchi M, Ichinose H. Patch Clamp Analysis of Voltage-Gated Na+ Currents in Human Retinal Bipolar Cells. Invest Ophthalmol Vis Sci 2006; 47: ARVO E-Abstr. 3750.

29. Ohkuma M, Kawai F, Horiguchi M, Miyachi E. Patch-clamp recording of human retinal photoreceptors and bipolar cells. Photochem Photobiol 2007 Mar-Apr; 83:317-22.

30. Shingai R, Christensen BN. Excitable properties and voltage-sensitive ion conductances of horizontal cells isolated from catfish (Ictalurus punctatus) retina. J Neurophysiol 1986; 56:32-49.

31. Lasater EM. Ionic currents of cultured horizontal cells isolated from white perch retina. J Neurophysiol 1986; 55:499-513.

32. Ueda Y, Kaneko A, Kaneda M. Voltage-dependent ionic currents in solitary horizontal cells isolated from cat retina. J Neurophysiol 1992; 68:1143-50.

33. Lohrke S, Hofmann HD. Voltage-gated currents of rabbit A- and B-type horizontal cells in retinal monolayer cultures. Vis Neurosci 1994 Mar-Apr; 11:369-78.

34. Schubert T, Weiler R, Feigenspan A. Intracellular calcium is regulated by different pathways in horizontal cells of the mouse retina. J Neurophysiol 2006; 96:1278-92.

35. Kawai F, Horiguchi M, Suzuki H, Miyachi E. Na(+) action potentials in human photoreceptors. Neuron 2001; 30:451-8.

36. Kawai F, Horiguchi M, Ichinose H, Ohkuma M, Isobe R, Miyachi E. Suppression by an h current of spontaneous Na+ action potentials in human cone and rod photoreceptors. Invest Ophthalmol Vis Sci 2005; 46:390-7.

37. Rasband MN, Peles E, Trimmer JS, Levinson SR, Lux SE, Shrager P. Dependence of nodal sodium channel clustering on paranodal axoglial contact in the developing CNS. J Neurosci 1999; 19:7516-28.

38. Sherry DM, Mitchell R, Standifer KM, du Plessis B. Distribution of plasma membrane-associated syntaxins 1 through 4 indicates distinct trafficking functions in the synaptic layers of the mouse retina. BMC Neurosci 2006; 7:54.

39. Sherry DM, Wang MM, Bates J, Frishman LJ. Expression of vesicular glutamate transporter 1 in the mouse retina reveals temporal ordering in development of rod vs. cone and ON vs. OFF circuits. J Comp Neurol 2003; 465:480-98.

40. Sherry DM, Wang MM, Frishman LJ. Differential distribution of vesicle associated membrane protein isoforms in the mouse retina. Mol Vis 2003; 9:673-88 <http://www.molvis.org/molvis/v9/a81/>.

41. Shi SR, Cote RJ, Taylor CR. Antigen retrieval immunohistochemistry: past, present, and future. J Histochem Cytochem 1997; 45:327-43.

42. McGinnis S, Madden TL. BLAST: at the core of a powerful and diverse set of sequence analysis tools. Nucleic Acids Res 2004; 32:W20-5.

43. Dawson WW, Trick GL, Litzkow CA. Improved electrode for electroretinography. Invest Ophthalmol Vis Sci 1979; 18:988-91.

44. Saszik SM, Robson JG, Frishman LJ. The scotopic threshold response of the dark-adapted electroretinogram of the mouse. J Physiol 2002; 543:899-916.

45. Hughes A. A schematic eye for the rat. Vision Res 1979; 19:569-88.

46. Dureau P, Bonnel S, Menasche M, Dufier JL, Abitbol M. Quantitative analysis of intravitreal injections in the rat. Curr Eye Res 2001; 22:74-7.

47. Osorio N, Alcaraz G, Padilla F, Couraud F, Delmas P, Crest M. Differential targeting and functional specialization of sodium channels in cultured cerebellar granule cells. J Physiol 2005; 569:801-16.

48. Bartoo AC, Sprunger LK, Schneider DA. Expression and distribution of TTX-sensitive sodium channel alpha subunits in the enteric nervous system. J Comp Neurol 2005; 486:117-31.

49. Ma YP, Cui J, Pan ZH. Heterogeneous expression of voltage-dependent Na+ and K+ channels in mammalian retinal bipolar cells. Vis Neurosci 2005 Mar-Apr; 22:119-33.

50. Liu IS, Chen JD, Ploder L, Vidgen D, van der Kooy D, Kalnins VI, McInnes RR. Developmental expression of a novel murine homeobox gene (Chx10): evidence for roles in determination of the neuroretina and inner nuclear layer. Neuron 1994; 13:377-93.

51. Ohno S, Kawasaki H, Imajoh S, Suzuki K, Inagaki M, Yokokura H, Sakoh T, Hidaka H. Tissue-specific expression of three distinct types of rabbit protein kinase C. Nature 1987 Jan 8-14; 325:161-6.

52. Rohrenbeck J, Wassle H, Heizmann CW. Immunocytochemical labelling of horizontal cells in mammalian retina using antibodies against calcium-binding proteins. Neurosci Lett 1987; 77:255-60.

53. Pasteels B, Rogers J, Blachier F, Pochet R. Calbindin and calretinin localization in retina from different species. Vis Neurosci 1990; 5:1-16.

54. Peichl L, Gonzalez-Soriano J. Morphological types of horizontal cell in rodent retinae: a comparison of rat, mouse, gerbil, and guinea pig. Vis Neurosci 1994 May-Jun; 11:501-17.

55. Johnson J, Tian N, Caywood MS, Reimer RJ, Edwards RH, Copenhagen DR. Vesicular neurotransmitter transporter expression in developing postnatal rodent retina: GABA and glycine precede glutamate. J Neurosci 2003; 23:518-29.

56. Nakanishi S, Masu M, Bessho Y, Nakajima Y, Hayashi Y, Shigemoto R. Molecular diversity of glutamate receptors and their physiological functions. EXS 1994; 71:71-80.

57. Bui BV, Fortune B. Ganglion cell contributions to the rat full-field electroretinogram. J Physiol 2004; 555:153-73.

58. Toda K, Bush RA, Humphries P, Sieving PA. The electroretinogram of the rhodopsin knockout mouse. Vis Neurosci 1999 Mar-Apr; 16:391-8.

59. Naarendorp F, Sato Y, Cajdric A, Hubbard NP. Absolute and relative sensitivity of the scotopic system of rat: electroretinography and behavior. Vis Neurosci 2001 Jul-Aug; 18:641-56.

60. Robson JG, Maeda H, Saszik SM, Frishman LJ. In vivo studies of signaling in rod pathways of the mouse using the electroretinogram. Vision Res 2004; 44:3253-68.

61. Robson JG, Saszik SM, Ahmed J, Frishman LJ. Rod and cone contributions to the a-wave of the electroretinogram of the macaque. J Physiol 2003; 547:509-30.

62. Lyubarsky AL, Falsini B, Pennesi ME, Valentini P, Pugh EN Jr. UV- and midwave-sensitive cone-driven retinal responses of the mouse: a possible phenotype for coexpression of cone photopigments. J Neurosci 1999; 19:442-55.

63. Akula JD, Lyubarsky AL, Naarendorp F. The sensitivity and spectral identity of the cones driving the b-wave of the rat electroretinogram. Vis Neurosci 2003 Mar-Apr; 20:109-17.

64. Nusinowitz S, Hood DC, Birch DG. Rod transduction parameters from the a wave of local receptor populations. J Opt Soc Am A Opt Image Sci Vis 1995; 12:2259-66.

65. Friedburg C, Thomas MM, Lamb TD. Time course of the flash response of dark- and light-adapted human rod photoreceptors derived from the electroretinogram. J Physiol 2001; 534:217-42.

66. Mojumder DK, Frishman LJ. TTX-blockade of spiking activity in the rat retina reduces the amplitude of the ERG b-wave originating from cone bipolar cells. Invest Ophthalmol Vis Sci 2005; 46: ARVO E-Abstr. 2252.

67. Bleakman D, Lodge D. Neuropharmacology of AMPA and kainate receptors. Neuropharmacology 1998 Oct-Nov; 37:1187-204.

68. Miguel-Hidalgo JJ, Snider CJ, Angelides KJ, Chalupa LM. Voltage-dependent sodium channel alpha subunit immunoreactivity is expressed by distinct cell types of the cat and monkey retina. Vis Neurosci 1994 Mar-Apr; 11:219-28.

69. Brysch W, Creutzfeldt OD, Luno K, Schlingensiepen R, Schlingensiepen KH. Regional and temporal expression of sodium channel messenger RNAs in the rat brain during development. Exp Brain Res 1991; 86:562-7.

70. Black JA, Yokoyama S, Higashida H, Ransom BR, Waxman SG. Sodium channel mRNAs I, II and III in the CNS: cell-specific expression. Brain Res Mol Brain Res 1994; 22:275-89.

71. Felts PA, Yokoyama S, Dib-Hajj S, Black JA, Waxman SG. Sodium channel alpha-subunit mRNAs I, II, III, NaG, Na6 and hNE (PN1): different expression patterns in developing rat nervous system. Brain Res Mol Brain Res 1997; 45:71-82.

72. Fisher SK, Boycott BB. Synaptic connections made by horizontal cells within the outer plexiform layer of the retina of the cat and the rabbit. Proc R Soc Lond B Biol Sci 1974; 186:317-31.

73. Kolb H, Normann RA. A-type horizontal cells of the superior edge of the linear visual streak of the rabbit retina have oriented, elongated dendritic trees. Vision Res 1982; 22:905-16.

74. Pinchuk GV, Malysheva MK, Pinchuk LN, Gerasymenko OV, Zhukareva VA. Possible relationship of brain cytoplasmic tetrodotoxin-sensitive protein to voltage-gated sodium channel shown by monoclonal antibody. J Neuroimmunol 1990; 26:91-6.

75. Gong B, Rhodes KJ, Bekele-Arcuri Z, Trimmer JS. Type I and type II Na(+) channel alpha-subunit polypeptides exhibit distinct spatial and temporal patterning, and association with auxiliary subunits in rat brain. J Comp Neurol 1999; 412:342-52.

76. Schmidt J, Rossie S, Catterall WA. A large intracellular pool of inactive Na channel alpha subunits in developing rat brain. Proc Natl Acad Sci U S A 1985; 82:4847-51.

77. Sieving PA, Murayama K, Naarendorp F. Push-pull model of the primate photopic electroretinogram: a role for hyperpolarizing neurons in shaping the b-wave. Vis Neurosci 1994 May-Jun; 11:519-32.

78. Viswanathan S, Frishman LJ, Robson JG, Harwerth RS, Smith EL 3rd. The photopic negative response of the macaque electroretinogram: reduction by experimental glaucoma. Invest Ophthalmol Vis Sci 1999; 40:1124-36.

79. Frishman LJ. Electrogenesis of the ERG. In: Hinton DR, editor. Retina. Vol 1 Basic Science and Inherited Retinal Diseases. 4 ed: Elsevier Mosby; 2005. p. 103-35.

80. Copenhagen D. Is the retina going digital? Neuron 2001; 30:303-5.

81. Massey SC, Miller RF. Excitatory amino acid receptors of rod- and cone-driven horizontal cells in the rabbit retina. J Neurophysiol 1987; 57:645-59.

82. Aoyama T, Kamiyama Y, Usui S, Blanco R, Vaquero CF, de la Villa P. Ionic current model of rabbit retinal horizontal cell. Neurosci Res 2000; 37:141-51.

83. Blanco R, Vaquero CF, de la Villa P. Action potentials in axonless horizontal cells isolated from the rabbit retina. Neurosci Lett 1996; 203:57-60.

84. Cook EP, Johnston D. Voltage-dependent properties of dendrites that eliminate location-dependent variability of synaptic input. J Neurophysiol 1999; 81:535-43.

85. Ichinose T, Lukasiewicz PD. Ambient light regulates sodium channel activity to dynamically control retinal signaling. J Neurosci 2007; 27:4756-64.

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