The G369i mutation results in an insertion of a glycine residue in a transmembrane domain, which prevents Ca²⁺ permeation through Ca_v1.4 (Maddox et al., 2020). The presence of rod synapses in G369i KI mice but not Ca_v1.4 KO mice indicates that the Ca_v1.4 protein and not its Ca²⁺ conductance is required for rod synapse assembly (Maddox et al., 2020). To test if this is also true for cones, we performed immunofluorescence and confocal analyses using antibodies against cone arrestin (CAR) and CtBP2 to label cone terminals (i.e. pedicles) and ribbons, respectively (Figure 1). In Ca_v1.4 KO mouse retinas, cone pedicles were shrunken and retracted into the outer nuclear layer (ONL, Figure 1a) with little evidence of elongated ribbons (Figure 1b). In contrast, cone pedicles in G369i KI mice were normally localized in the outer plexiform layer (OPL, Figure 1a) and were populated by multiple ribbons that were associated with labeling for Ca_v1.4 (Figure 1b). Some abnormalities were apparent in G369i KI cone pedicles, such as extremely elongated ribbons and telodendria extending toward the inner nuclear layer (Figure 1b). These results indicate that Ca_v1.4 Ca²⁺ signals are dispensable for the integrity of ribbons and the pedicle but are necessary for their overall refinement.

Figure 1

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A prevailing yet unsupported hypothesis regarding the relatively mild visual phenotypes in CSNB2 is that additional Ca_v subtypes may compensate for Ca_v1.4 loss-of-function in cones. If so, then Ca²⁺ currents (I_Ca) mediated by these subtypes should be evident in cones of Ca_v1.4 KO and G369i KI mice. To test this, we performed patch clamp recordings of cones in retinal slices of adult WT, G369i KI, and Ca_v1.4 KO mice under conditions designed to isolate I_Ca. During voltage ramps in WT cones, I_Ca activated around –50 mV and peaked near –20 mV, consistent with the properties of Ca_v1.4 (Figure 2a–d). While not observed in rods of G369i KI or Ca_v1.4 KO mice (Maddox et al., 2020), a small-amplitude I_Ca that activated around –60 mV and peaked near –35 mV was detected in cones of these mice (Figure 2a–d). To uncover this low voltage-activated I_Ca, we used a faster voltage ramp (0.5 mV/ms) than in the previous study of G369i KI rods (~0.15 mV/ms) Maddox et al., 2020. With this faster voltage ramp, I_Ca was not apparent in G369i KI rods (Figure 2—figure supplement 1). Therefore, the aberrant I_Ca is an adaptation to Ca_v1.4 loss-of-function in cones but not rods.

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Values obtained from recordings that were used for analyses in Figure 2C, D, F, H, I .

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We next used step depolarizations to identify the type of Ca_v channel underlying I_Ca in G369i KI and Ca_v1.4 KO cones. Analysis of current-voltage (I-V) and conductance-voltage (G-V) relationships revealed a significant, hyperpolarizing shift (~10 mV) in the voltage of half-maximal activation (V_h) in G369i KI cones (Figure 2c and d; Table 1). I_Ca in G369i KI cones was not sustained as in WT cones but inactivated rapidly during 500 ms step depolarizations (Figure 2e). For G369i KI cones, the overlay of the conductance-voltage (Figure 2d) and inactivation curves revealed a sizeable window current (Figure 2f). These features of I_Ca in G369i KI and Ca_v1.4 KO cones resembled those of Ca_v3 T-type channels rather than Ca_v1.4 Perez-Reyes, 2003.

In a previous study, a Ca_v3-like current was detected in patch clamp recordings of cone pedicles in WT mouse cones (Davison et al., 2022). However, in recordings of WT mouse cone somas, we did not observe a low-voltage activated component in the I-V or G-V curves that would be indicative of a Ca_v3 subtype (Figure 2c and d). To further test for a Ca_v3 contribution to I_Ca in WT mouse cones, we used a double pulse voltage protocol where I_Ca was evoked before (P1) and after (P2) a 500 ms step to –60 mV (Figure 2g). With this protocol, the inactivation of Ca_v3 channels by the –60 mV step should lead to a reduction in the amplitude of the P2 vs the P1 current. While such a reduction was clearly evident in cones of G369i KI mice (i.e. P2/P1 <1), there was no significant difference in the P2 and P1 currents in cones of WT mice (Figure 2g–i).

We also used antagonists selective for Ca_v1 and Ca_v3 channels (isradipine and ML 218, respectively). As expected, I_Ca in WT cones was significantly but incompletely suppressed by isradipine (Figure 3a and b). The V_h of the I-V was similar before (–36.5±0.8 mV) and after (–38.3±1.8 mV) perfusion of isradipine (t(3,4)=0.90, p=0.43 by paired t-test), suggesting that the residual I_Ca after exposure to isradipine arose from unblocked Ca_v1.4 rather than Ca_v3 channels. While ML 218 modestly inhibited I_Ca in some WT cones, the effect was not statistically significant (Figure 3c and d) and could be attributed to weak activity on Ca_v1.4. In transfected HEK293T cells, ML 218 modestly inhibited I_Ca and caused a negative shift in the V_h for Ca_v1.4, in contrast to its strong inhibition of Ca_v3.2 (Figure 3—figure supplement 1). Like its effects on Ca_v3.2, ML 218 nearly abolished I_Ca in cones of G369i KI mice whereas isradipine had little effect (Figure 3a–d).

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Values obtained from recordings that were used for analyses in Figure 3b and d.

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Finally, we recorded from ground squirrel and macaque cones, where the large amplitude of I_Ca facilitates pharmacological and biophysical analyses. Consistent with its actions on Ca_v1.4 in WT mouse cones (Figure 3b) and transfected HEK293T cells (Figure 3—figure supplement 1), ML 218 caused an insignificant inhibition of peak I_Ca (–7.0 ± 20.8%, n=6 cones) as well as a negative shift in the voltage-dependence of activation in ground squirrel cones (ΔV_1/2 = -1.16 ± 0.94 mV, n=6, mean ± SD, p=0.029, t-test; Figure 4a and b). By contrast, the application of isradipine dramatically suppressed I_Ca in cones in various regions of the ground squirrel retina (Figure 4c and d). Subsequent application of ML 218 did not reveal a Ca_v3-like current: the I-V and G-V of the isradipine + ML 218-sensitive current were shifted in the positive rather than the negative direction relative to the isradipine-sensitive current (Figure 4c–g). This result is consistent with the time- and voltage-dependent block of Ca_v1 channels by dihydropyridine antagonists such as isradipine (Koschak et al., 2003). As a positive control, we confirmed that ML 218 blocked a prominent Ca_v3-type current in ground squirrel type 3 a OFF cone bipolar cells (Figure 4h and i). In recordings of macaque cone pedicles, we compared the properties of I_Ca using a holding voltage of –90 mV or –50 mV, the latter of which should inactivate Ca_v3 channels (Figure 5a). There was no difference in the I-V or G-V relationships obtained at these holding voltages (Figure 5b–e). As in WT mouse cones (Figure 2g–i), there was also no difference in I_Ca recorded before and after an inactivating pulse to –60 mV (Figure 5f and g). We conclude that Ca_v3 channels do not normally contribute to I_Ca in cones of WT mice, ground squirrels, and macaques.

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Representative traces and values were obtained from recordings that were used for analyses in Figure 4.

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Values obtained from recordings that were used for analyses in Figure 5b–e and g.

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To test whether the appearance of the Ca_v3-mediated I_Ca in G369i KI cones could result from an increase in the expression of a specific Ca_v3 subtype (i.e. Ca_v3.1, Ca_v3.2, or Ca_v3.3), we initially tried labeling with commercially available antibodies against Ca_v3.2. However, the data were unreliable since these antibodies produced a similar labeling pattern in Ca_v3.2 KO retinal tissue (data not shown). As an alternative, we performed single-cell RNA sequencing on cells isolated from the retina of WT and G369i KI mice (Figure 3—figure supplement 2). We sequenced 28,782 cells and, with unsupervised clustering at resolution = 0.5, we grouped the cells into 20 clusters. Clusters 8 and 16 were identified as cones based on the expression of cone marker genes (e.g. Opn1sw, Opn1mw, Arr3, Gnat2). Consistent with previous studies (Davison et al., 2022; Williams et al., 2022), Cacna1h encoding the Ca_v3.2 subtype was the major Ca_v3 transcript expressed in WT mouse cones. There was no significant difference in Cacna1h expression between WT and G369i KI cones according to this analysis (Figure 3—figure supplement 2). Thus, the enhanced activity of Ca_v3 channels in cones of G369i KI mice results from a mechanism other than increased transcription of Cacna1h.

The replacement of Ca_v1.4 with Ca_v3 as the primary conduit for Ca²⁺ influx in G369i KI cones allowed us to dissect the importance of Ca_v1.4 Ca²⁺ signals for the molecular and structural organization of the cone synapse. To this end, we first analyzed WT and G369i KI cone synapses by immunofluorescence and confocal microscopy. Presynaptic proteins such as bassoon and members of the postsynaptic signaling complex in depolarizing (ON) cone bipolar cells (GPR179, mGluR6, and TRPM1 Martemyanov and Sampath, 2017) were enriched near cone ribbons in G369i KI mice as in WT mice (Figure 6a). Compared to WT mice, the labeled structures in G369i KI mice occupied a larger volume (Figure 6b and c) but were of lower intensity (Figure 6d), suggesting an increase in the spread rather than in the levels of synaptic proteins. Unlike in WT mice, the volume of presynaptic and postsynaptic proteins increased linearly with the volume of the pedicle in G369i KI mice (Figure 6e–i). These results show that the molecular determinants of the cone synapse can assemble in G369i KI mice but change in their sub-synaptic distribution in ways that correlate with the expansion of the cone pedicle.

Figure 6

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Values obtained from images that were used for analyses in Figure 6b–i.

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The cone synapse is structurally complex, with the dendritic tips of two horizontal cells and an intervening ON cone bipolar cell invaginating deeply into the pedicle near the ribbon (Haverkamp et al., 2000). To test how the switch in Ca_v subtypes might affect this arrangement of postsynaptic partners, we generated 3D reconstructions of cone synapses by serial block-face scanning electron microscopy (SBFSEM; Figure 7). As with the enlarged synaptic contacts (Figure 6), structural modifications in G369i KI pedicles were evident. Compared to WT, ribbons in G369i KI pedicles appeared disorganized and were often parallel rather than perpendicular to the presynaptic membrane (Figure 7a–c). Consistent with our confocal analyses (Figure 1), G369i KI cone pedicles extended telodendria in multiple directions rather than just apically (Figure 7a). In addition, a slightly larger fraction of synaptic sites in the G369i KI pedicle (35% vs 14% in WT) formed incorrect postsynaptic partnerships including with glia (Figure 7b–d’; Table 2). Nevertheless, the number of ribbons is normal in G369i KI cones, and about half of the ribbons make invaginating contacts with the appropriate cell types (i.e. both horizontal cells and cone bipolar cells; Table 2). On average, there were more vesicles associated with ribbons in cones of G369i KI mice (245 per ribbon) than of WT mice (178 per ribbon; Table 2), but the difference did not reach statistical significance. Together with our immunofluorescence data, these results indicate that cone synapses undergo relatively subtle morphological changes in G369i KI mice, perhaps in response to the loss of Ca_v1.4 Ca²⁺ signals.

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Compared to Ca_v1.4 KO mice, the relative preservation of cone synapses in G369i KI mice allowed the unique opportunity to test whether a Ca_v subtype other than Ca_v1.4 could support synaptic release. To this end, we analyzed synaptic transmission between cones and horizontal cells (HCs). In the darkness, glutamate released from cones depolarizes horizontal cells via activation of α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic (AMPA)/kainate receptors (Hack et al., 2001; Brandstätter et al., 1997). The resulting excitatory postsynaptic current (EPSC) undergoes a decline in response to light stimuli that hyperpolarize cones (Feigenspan and Babai, 2015). In WT HCs, a 1 s light flash (λ=410 nm) inhibited the standing EPSC, which is reflected as an outward (hyperpolarizing ‘ON’) current (Figure 8a). Upon termination of the light, an inward (depolarizing ‘OFF’) current signaled the resumption of the EPSC (Figure 8a). Both the ON and OFF responses in WT HCs increased with light intensity, reflecting the impact of luminance on the presynaptic membrane potential of cones and the subsequent change in glutamate release from their terminals (Figure 8b).

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Representative traces and values were obtained from recordings that were used for analyses in Figure 8.

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Consistent with the lack of mature ribbons and abnormal cone pedicles (Figure 1), HC light responses were negligible in Ca_v1.4 KO mice (Figure 8a and b). In contrast, the ON and OFF responses were present in G369i KI HCs although significantly lower in amplitude than in WT HCs (Figure 8a and b). As expected, the EPSC in darkness and the light responses were abolished by the AMPA/kainate receptor antagonist, DNQX, in both WT and G369i KI HCs (Figure 8c and d). In addition to its smaller amplitude, the transient nature of the ON response in G369i KI HCs suggested inadequate cessation of cone glutamate release by light (Figure 8b). Slow deactivation of Ca_v3 channels and/or their activation at negative voltages (Perez-Reyes, 2003) could give rise to Ca²⁺ signals that support release following light-induced hyperpolarization of G369i KI cones.

To test if the diminished HC light responses correlated with lower presynaptic Ca²⁺ signals in G369i KI cones, we performed 2-photon imaging of vertical slices prepared from the whole retina that was incubated with the Ca²⁺ indicator Fluo3-AM and Alexa-568-conjugated peanut agglutinin (PNA) to demarcate regions of interest (ROIs) corresponding to cone pedicles. With this approach, Fluo3 fluorescence was detected only in photoreceptors and ganglion cells and not inner retinal cell-types (e.g. horizontal cells, bipolar cells, Mueller cell soma). Thus, Ca²⁺ signals reported by Fluo3 fluorescence near PNA-labeling originated primarily from cones. In WT cones, depolarization by K⁺ (50 mM) caused a sustained increase of Ca²⁺ (ΔF/F₀=1.85 ± 0.1, n=11 terminals) that was significantly lower after the addition of isradipine (ΔF/F₀=1.44 ± 0.07, p=0.0015 by repeated measures one-way ANOVA with Tukey’s multiple comparison posthoc test; F[1.608, 16.08]=55.17; p<0.0001; Figure 9a and b). The Ca²⁺ signal remaining in the presence of isradipine could arise from unblocked Ca_v1.4 channels and/or Ca²⁺ release from intracellular stores (Križaj, 2012). In G369i KI cones, the peak amplitude of the K⁺-evoked Ca²⁺ signal (ΔF/F₀=1.07 ± 0.02, n=14 terminals) was significantly lower than in WT cones (t(23)=8.74, p<0.0001 by unpaired t-test, n=14 terminals). The decrease in the Ca²⁺ signal in the G369i KI cone pedicles during isradipine exposure paralleled that during the depolarization (Figure 9a), suggesting it was not a consequence of Ca_v1 blockade. The lower amplitude and more transient Ca²⁺ signals mediated by Ca_v3 channels may be cleared more rapidly in G369i KI cones as compared to prolonged Ca_v1.4-initiated Ca²⁺ signals in WT cones. Taken together, our results suggest that Ca_v3 channels nominally support Ca²⁺ signals and synaptic transmission in cones of G369i KI mice.

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Values obtained from time series ROIs that were used for analyses in Figure 9b.

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While HCs play important roles in lateral inhibition of photoreceptor output, the vertical dissemination of visual information from cones to the inner retina is relayed by glutamate to ON and OFF cone bipolar cells (CBCs) which express mGluR6 and AMPA/kainate receptors, respectively. To test for the existence of cone-to-CBC transmission in G369i KI mice, we recorded electroretinograms (ERGs) under light-adapted conditions using Ca_v1.4 KO mice as a negative control (Figure 10a–d). In these recordings, the light-induced response of photoreceptors and the postsynaptic response of ON CBCs correspond to the a- and b-waves, respectively. Unlike in Ca_v1.4 KO mice, the a-waves of G369i KI mice resembled those in WT mice, which indicates that cones do not degenerate in this mouse strain (Figure 10a and b). While greatly reduced in amplitude, the b-wave was measurable in G369i KI mice and significantly larger than in Ca_v1.4 KO mice at the highest light intensities (F(14,104)=41, p<0.0001, post-hoc Tukey’s p=0.0038 at 2.3 log cd·s/m²; Figure 10a and b). We also recorded flicker ERGs using 10 Hz light stimuli that can isolate cone pathways involving both ON and OFF CBCs (Nusinowitz et al., 2007; Vinberg et al., 2015). In WT mice, flicker responses exhibited two peaks, one at a lower irradiance (–2 log cd·s/m²) and one at higher irradiance (0.5 log cd·s/m²; Figure 10c and d). The peak at the lower irradiance is attributed to responses in both rod and cone pathways, and the peak at a higher irradiance is attributed to responses exclusively in cone pathways (Nusinowitz et al., 2007; Vinberg et al., 2015). Flicker responses in G369i KI mice showed a similar non-monotonic relation as in WT mice but were significantly lower in amplitude (F(26,143)=51.18, p<0.0001, post-hoc Tukey’s p=0.0237 at 2.5 log cd·s/m²; Figure 10c and d). The inverted flicker responses at higher illuminations in G369i KI mice were absent in Ca_v1.4 KO mice and may result from the hyperpolarizing contribution of cone-to-OFF CBC transmission (Figure 10c).

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Representative electroretinogram (ERG) traces and values obtained from ERGs and behavioral swim tests that were used for analyses in Figure 10.

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To test whether the cone synaptic responses in G369i KI mice enable vision-guided behavior, we used a swim test that assesses the ability of mice to identify a visible platform (Bimonte-Nelson et al., 2015). Under scotopic (dark) conditions, G369i KI and Ca_v1.4 KO mice took significantly longer to find the platform compared to WT mice (Figure 10e and f). These results are consistent with flicker response assays (Figure 10c and d) as well as the absence of I_Ca in rods and rod-to-rod bipolar cell synaptic transmission in G369i KI mice (Maddox et al., 2020). Under photopic (light) conditions, G369i KI mice but not Ca_v1.4 KO mice performed as well as WT mice (Figure 10e and f). Thus, G369i KI mice have enough visual function under photopic conditions to quickly find the platform. Collectively, our results suggest that Ca_v3 channels can support cone synaptic responses that are sufficient for visual behavior in G369i KI but not Ca_v1.4 KO mice.

As shown for rod synapses (Maddox et al., 2020), ribbons and other components of the pre- and post-synaptic complex assemble at cone synapses in G369i KI mice (Figures 1, 6 and 7). Ca_v3 Ca²⁺ signals are dispensable for this process since their presence in Ca_v1.4 KO cones is not accompanied by any semblance of ribbon synapses (Figure 1b). An intimate relationship between Ca_v1.4 and ribbons is supported by the colocalization of Ca_v1.4 and puncta resembling ribbon precursor spheres in the developing OPL (Liu et al., 2013). Ca_v1.4 interacts directly or indirectly with a variety of ribbon synapse-associated proteins including RIM (Kiyonaka et al., 2007), CAST/ERC2 (Kiyonaka et al., 2012), bassoon (tom Dieck et al., 2005), unc119 Alpadi et al., 2008; Haeseleer, 2008; Haeseleer et al., 2004, and a major component of the ribbon, RIBEYE (Schmitz et al., 2000). Ca_v1.4 could pioneer sites of ribbon assembly perhaps by serving as a docking or nucleation site for the active zone. At the same time, ribbon-associated proteins help cluster Ca_v1.4 near ribbons since knockout of these proteins in mice leads to fewer presynaptic Ca_v1.4 channels and in some cases shortened ribbons (tom Dieck et al., 2012). Regardless of the mechanism, our findings show that the formation of photoreceptor synaptic complexes does not require Ca²⁺ influx through Ca_v1.4 channels.

Our drop-seq analysis aligns with previous studies indicating that Cacna1h is expressed at extremely low levels compared to Cacna1f in WT mouse cones (Figure 3—figure supplement 2; Williams et al., 2022; Macosko et al., 2015). However, in contrast to recordings of WT mouse cone pedicles in a previous study (Davison et al., 2022), we found no evidence for Ca_v3-mediated currents in somatic recordings of cones in WT mice (Figures 2 and 3). We do not feel the discrepancy results from the different recording configurations given the electrically compact nature of cones. Moreover, recordings from macaque cone pedicles and from the ellipsoids of cylindrically shaped ground squirrel cones also did not reveal a Ca_v3-mediated current (Figures 4 and 5). We acknowledge that there are caveats of the pharmacological approach used in our study as well as the previous work (Davison et al., 2022). For example, dihydropyridine antagonists such as isradipine have relatively low affinity for some Ca_v1 subtypes, including Ca_v1.4 Koschak et al., 2003; Koschak et al., 2001, and have strongly voltage-dependent blocking properties. At a holding voltage of –90 mV, isradipine at 1 μM causes only ~80% inhibition of Ca_v1.4 with a greater block at depolarized voltages (Koschak et al., 2003), which agrees with our recordings of cones in WT mice (Figure 3a) and ground squirrels (Figure 4b–e). Moreover, in the low micromolar range, Ca_v3 blockers such as Z944 Tringham et al., 2012 and ML 218 (Figure 3—figure supplement 1) have nonspecific effects on Ca_v1 current amplitude and activation voltage. Thus, Ca_v channels mediating an I_Ca that is suppressed by Ca_v3 blockers and spared by dihydropyridines at negative voltages may be mistakenly categorized as Ca_v3 subtypes. In our experiments, the biophysical properties of the isradipine-sensitive I_Ca in cones of WT mice and ground squirrels resembled only those of Ca_v1, whereas those of the ML 218-sensitive I_Ca in cones of G369i KI mice resembled only those of Ca_v3. Therefore, we favor the interpretation that Ca_v3 contributes to I_Ca in mouse cones only upon silencing of Ca_v1.4 either by KO of the corresponding protein or by silencing its conductance.

Since Cacna1h expression was similar in WT and G369i KI cones (Figure 3—figure supplement 2), what causes the increased Ca_v3 channel activity in the latter? Among the Ca_v3 subtypes, Ca_v3.2 is uniquely sensitive to post-translational modifications that facilitate the opening of the channel and/or its cell surface trafficking (Cai et al., 2021). For example, Ca_v3.2 interacts with the deubiquitinating enzyme USP5 which enhances levels of Ca_v3.2 protein levels by suppressing its proteasomal degradation (García-Caballero et al., 2014). In addition, Ca_v3.2 is phosphorylated by calmodulin-dependent kinase II Welsby et al., 2003 and cyclin-dependent kinase 5 (Gomez et al., 2020), which enhances channel activation and current density, respectively. Further interrogation of our drop-seq dataset could illuminate novel Ca_v3.2-regulatory pathways that are modified in G369i KI cones.

Despite having the same complement of proteins as in WT mice, cone synapses in G369i KI mice were enlarged and made errors in postsynaptic partner selection (Figures 6 and 7). While I-V curves indicate similar peak I_Ca amplitudes in cones of WT and G369i KI mice near the membrane potential of cones in darkness (−45 to –50 mV Ingram et al., 2019, Figure 2a and c), the strong inactivation of Ca_v3 channels and/or reduced Ca²⁺ release from intracellular stores likely account for the greatly diminished Ca²⁺ signals in G369i KI pedicles (Figure 9). Paradoxically, reductions in presynaptic Ca²⁺ in rod photoreceptors correlated with illumination-dependent shrinkage of ribbons in vivo and in vitro (Dembla et al., 2020; Spiwoks-Becker et al., 2004; Regus-Leidig et al., 2010). However, ribbon synapse size increases with low presynaptic Ca²⁺ in zebrafish sensory hair cells Sheets et al., 2012. The mechanism involves decreased mitochondrial Ca²⁺ uptake and an increase in the redox state of nicotinamide adenine dinucleotide (NAD⁺/NADH ratio) (Wong et al., 2019). Ca_v3 Ca²⁺ signals may decay too quickly in G369i KI cones (Figure 9) to enable mitochondrial Ca²⁺ uptake or other mechanisms that trim ribbons.

An intriguing result was the linear increase in ribbon synapse size with pedicle volume in G369i KI but not in WT mice (Figure 6e–i), which could represent a form of homeostatic synaptic scaling as has been demonstrated at the neuromuscular junction (Goel et al., 2019; Hong et al., 2020). Similarly, the presence of some non-invaginating contacts with incorrect partner pairings in G369i KI mice (Figure 7, Table 2) could be an attempt to increase synaptic connectivity to compensate for weakened synaptic output. Consistent with this possibility, diminished presynaptic Ca²⁺ signals and non-invaginating cone synapses also characterize the retina of mice lacking the extracellular α₂δ−4 subunit of Ca_v1.4 (Kerov et al., 2018). The cone synapses of G369i KI mice and α₂δ−4 KO mice are clearly functional given that photopic visual behavior is preserved in both mouse strains Figure 10e and f; Kerov et al., 2018 and, in the case of G369i KI mice, HCs exhibit EPSCs whose amplitude decreases with light and AMPA/kainate receptor blockade (Figure 8). Defects in cone synaptic transmission that correlate with aberrant cone synapse morphology and connectivity also typify a mouse strain with a gain-of-function mutation in Ca_v1.4 linked to CSNB2 (Zanetti et al., 2021; Knoflach et al., 2013). Thus, balanced presynaptic Ca²⁺ signals may be needed to encode the correct post-synaptic wiring and structure of the cone synapse.

In addition to structural abnormalities in cone synapses, the inability of Ca_v3 channels to support sustained presynaptic Ca²⁺ entry likely contributed to the impaired light responses in HCs and CBCs in G369i KI mice (Figures 8 and 10). Due to their slow activation and strong inactivation (Perez-Reyes et al., 2009), Ca_v3 channels are not expected to fuel the Ca²⁺ nanodomains that support fast and sustained components of release, both of which occur only at ribbon sites in cones (Snellman et al., 2011; Choi et al., 2008). Ca_v3 channels may also be located further from the ribbon than Ca_v1.4, thus lowering the efficiency of coupling to exocytosis. Regardless of the features that differentiate the contributions of Ca_v3 and Ca_v1.4 to cone synaptic release, our findings provide further support for the widely held view that Ca_v1 channels are well-suited to support transmission at ribbon-type synapses.

Based on extremely heterogeneous clinical presentations, CSNB2 manifests as a spectrum of visual disorders that originate from various mutations in CACNA1F (Boycott et al., 2000; Bijveld et al., 2013). Even though some CSNB2 mutant Ca_v1.4 channels may traffic normally to the plasma membrane, many of these mutations are expected to produce non-functional, non-conducting Ca_v1.4 channels (Peloquin et al., 2007; Hoda et al., 2005). Yet, the visual phenotypes of CSNB2 patients are not as severe as the complete blindness in Ca_v1.4 KO mice, which lack any Ca_v1.4 protein expression and exhibit no signs of visual behavior (Figure 10e and f; Kerov et al., 2018). Collectively, our results suggest that G369i KI mice accurately model Ca_v1.4 channelopathies in CSNB2 patients that are characterized by a greater impairment in rod than in cone pathways. This interpretation is supported by our findings that G369i KI mice exhibit horizontal cell responses to bright but not dim illumination (Figure 8), ERG responses under conditions of light adaptation (Figure 10a–d) but not dark-adaptation (Maddox et al., 2020), and visual behavior under photopic but not scotopic conditions (Figure 10e and f). Together with the enlargement of synaptic sites, modest levels of synaptic release from cones of G369i KI mice may support the nominal transmission of visual information through cone pathways. G369i KI mice could also exhibit homeostatic alterations in the inner retina, which are known to support visual function when photoreceptor input is severely compromised (Shen et al., 2020; Leinonen et al., 2020; Care et al., 2020). Future studies of the retinal circuitry and visual behavior of G369i KI mice could identify compensatory pathways that are recruited upon Ca_v1.4 loss-of-function and how they might be targeted in novel therapies for CSNB2 and related disorders.

Values represent the median (25^th, 75^th quartiles). V_h and k were determined from Boltzmann fits of the G-V and steady-state inactivation curves. Time constant (tau) for activation was obtained from the exponential fit of the rising phase of I_Ca evoked by a 50 ms test pulse to a voltage near the peak of the I-V. Tau deactivation was determined from the exponential fit of the decay of the tail current evoked by repolarization to –90 mV from +20 mV. C_M, membrane capacitance; R_M, input resistance. p-values were determined by the Kruskal-Wallis test. ^a, relative to WT. ^b, relative to G369i KI.

Results represent the analysis of n=2 pedicles reconstructed by serial block face scanning electron microscopy in images obtained from WT or G369i KI mice (N=1 animal each). The first row of vesicles encircling and immediately apposed to the ribbon were quantified for all the z-planes of the ribbon. Sites were counted as 'invaginating' if postsynaptic partners were enveloped on all sides by the cone terminal for at least a couple of consecutive image planes.

All animal experiments were performed in accordance with guidelines approved by the National Institutes of Health and the Institutional Animal Care and Use Committees at the University of Texas at Austin and Northwestern University were approved by the Institutional Animal Care and Use Committee (AUP-2023–00170). The G369i KI Maddox et al., 2020 and Ca_v1.4 KO mouse strains were bred on the C57BL/6 J background strain for at least 10 generations. Adult male and female mice were used (6–12 weeks old), and aged-matched C57BL6/J mice were used as the control (WT) animals.

Mice between 6–8 weeks of age were anesthetized using isoflurane and euthanized by cervical dislocation. Eyes were enucleated and hemisected. The eye cups with retina were fixed on ice in 4% paraformaldehyde in 0.1 M phosphate buffer (PB) for 30 min. Fixed eye cups were then washed three times with 0.1 M PB containing 1% glycine followed by an infusion of 30% sucrose at 4 °C overnight. The eye cups were orientated along their dorsal-ventral axis and frozen in a 1:1 (wt/vol) mixture of Optimal Cutting Temperature compound and 30% sucrose in a dry ice/isopentane bath. Eye cups were cryosectioned at 20 µm on a Leica CM1850 cryostat (Leica Microsystems), mounted on Superfrost plus Micro Slides (VWR), dried for 5–10 min at 42 °C, and stored at –20 °C until used. Slides with mounted cryosections were warmed to room temperature, washed with 0.1 M PB for 30 min to remove the OCT/sucrose mixture and blocked with dilution solution (DS, 0.1 M PB/10% goat serum/0.5% Triton-X100) for 15 min or overnight at room temperature. All remaining steps were carried out at room temperature. All primary antibodies and appropriate secondary antibodies were diluted in DS at concentrations specified in the Key Resources Table. Sections were incubated with primary antibodies for 1 hr or overnight and then washed five times with 0.1 M PB. Sections were then incubated with secondary antibodies for 30 min and then washed five times with 0.1 M PB. Trace 0.1 M PB was removed, and sections were then mounted with #1.5 H coverslips (ThorLabs) using ProLong Glass Antifade Mountant with or without NucBlue (Thermo Fisher Scientific).

For double labeling with other rabbit polyclonal antibodies, CAR antibodies were conjugated with the CF647 fluorophore (CAR-647) using the Mix-n-Stain antibody labeling kit according to the manufacturer’s protocol (Biotium). Sections were processed first with rabbit polyclonal Ca_v1.4 or EAAT2 antibodies and corresponding secondary antibodies as described above. To prevent CAR-647 from binding to any available sites on the anti-rabbit secondary antibodies previously added to the sections, Ca_V1.4 or EAAT2 antibodies were readded to the sections and incubated for 30 min. After washing, the sections were incubated with CAR-647 for 1 hr, followed by wash steps and cover glass mounting.

Immunofluorescence in labeled retinal sections was visualized using an Olympus FV3000 confocal microscope (Tokyo, Japan) equipped with an UPlanApo 60 x oil HR objective (1.5 NA). Images were captured using the Olympus FLUOVIEW software package. Acquisition settings were optimized using a saturation mask to prevent signal saturation prior to collecting 16-bit. All confocal images presented are maximum z-projections. Images (256 × 256 pixels) used in analyses of synaptic proteins were collected using a 30 X optical zoom, 0.6 Airy disk aperture, and voxel size of 0.028 μm × 0.028 μm (X × Y × Z). Amira segmentation was used for generating 3D binary masks, and Amira 3D label analysis was used for quantification of immunofluorescent and masked images. Non-deconvolved images were used for all analyses. 3D binary masks of individual CAR-labeled pedicles were made by setting the threshold 1 standard deviation above the mean fluorescence intensity of each 3D Figure . and 3D binary masks of presynaptic labels (CtBP2, Ca_V1.4, and Bassoon) or postsynaptic labels (GPR179, mGluR6, and TRPM1) within or associated with the pedicle, respectively, were made by setting the threshold 3 standard deviations above the mean fluorescence intensity. To aid in presentation, high-mag images displayed in Figures 1b and 6a were deconvolved using cellSens software (Olympus), and any immunofluorescence corresponding to rod synaptic proteins was subtracted from the image. Experiments involving qualitative and/or quantitative analyses were performed independently at least three times, with one eye from per animal representing a biological replicate (three animals per genotype).

The cDNAs for Ca_V1.4 (GenBank: NM_019582), β_2X13 (GenBank: KJ789960), and α₂δ–4 (GenBank: NM_172364) were previously cloned into pcDNA3.1 Lee et al., 2015. The cDNA for Ca_V3.2 (GenBank: AF051946) was a gift from Dr. Edward Perez-Reyes, University of Virginia. All constructs were verified by DNA sequencing before use. Human embryonic kidney 293T (HEK293T) cells were cultured in Dulbecco’s Modified Eagle’s Medium with 10% FBS at 37 °C in 5% CO₂. At 70–80% confluence, the cells were co-transfected with cDNAs encoding human Ca_v1.4 (1.8μg) β_2X13 (0.6 μg), α₂δ–4 (0.6 μg), and enhanced GFP in pEGFP-C1 (Clonetech, 0.1 μg) or Ca_V3.2 (2μg) and pEGFP-C1 (0.1 μg) using FuGENE 6 transfection reagent according to the manufacturer’s protocol. Cells treated with the transfection mixture were incubated at 37 °C for 24 hr, dissociated using Trypsin-EDTA, and replated at a low density to isolate single cells. Replated cells were then incubated at 30 °C or 37 °C for an additional 24 hr before beginning experiments.

HEK293T extracellular recording solution contained the following (in mM): 140 Tris, 20 CaCl₂, 1 MgCl₂, pH 7.3 with methanesulfonic acid, and osmolarity 309 mOsm/kg. HEK293T internal recording solution contained the following: 140 NMDG, 10 HEPES, 2 MgCl₂, 2 Mg-ATP, 5 EGTA, pH 7.3 with methanesulfonic acid, osmolarity 358 mOsm/kg.

For recordings of I_Ca in mouse retina, extracellular recording solution contained the following (in mM): 115 NaCl, 2.5 KCl, 22.5 NaHCO₃, 1.25 NaH₂PO₄, 2 CaCl₂, 1 MgCl₂, 5 HEPES, 5 CsCl, 5.5 Glucose, osmolarity 290 mOsm/kg. Mouse cone intracellular solution contained the following (in mM): 105 CsMeSO₄, 20 TEA-Cl, 1 MgCl₂, 11 HEPES, 10 EGTA, 4 Mg-ATP, 10 phosphocreatine, 0.3 Na-GTP, pH 7.4 with CsOH, osmolarity 300 mOsm/kg.

For recordings of I_Ca in ground squirrel retina, extracellular recording solution contained the following (in mM): 10 HEPES, 85 NaCl, 3.1 KCl, 2.48 MgSO₄, 6 Glucose, 1 Na-succinate, 1 Na-malate, 1 Na-lactate, 1 Na-pyruvate, 2 CaCl₂, 25 NaHCO₃, and 20 TEA-Cl, osmolarity 285 ± 5 mOsm/kg. Intracellular solution contained the following (in mM): 80 CsCl, 10 BAPTA, 2 MgSO₄, 10 HEPES, 20 TEA-Cl, 5 Mg-ATP, and 0.5 Na-GTP, pH 7.35 with CsOH, osmolarity 285±5 mOsm/kg. For recordings of I_Aglu, extracellular recording solution contained (in mM): 125 NaCl, 3.0 KCl, 1.25 NaH₂PO₄, 25 NaHCO₃, 2 CaCl₂, 1 MgCl₂, 3 dextrose, 3 sodium pyruvate, 0.1 picrotoxin and 0.02 DNQX, and in indicated experiments 0.0013 TFB-TBOA. Intracellular I_Aglu contained (in mM): 125 KSCN, 10 TEA-Cl, 10 HEPES, 1 CaCl₂, 2 MgCl₂, 0.3 Na-GTP, 4 Mg-ATP, 10 K₂ phosphocreatine, 0.02 ZD7288. Mouse and ground squirrel extracellular slice recording solutions were equilibrated with 5% CO₂ /95% O₂ to a pH of ~7.5.

For recordings of light responses in horizontal cells of mouse retina, extracellular recording solution consisted of Ames’ media supplemented with 100 U/mL penicillin, 0.1 mg/mL streptomycin, and 22.6 mM NaHCO₃, osmolarity 280±5 mOsm/kg. The intracellular recording solution contained the following (in mM): 135 K-Aspartate, 10 KCl, 10 HEPES, 5 EDTA, 0.5 CaCl₂, 1 Mg-ATP, 0.2 Na-GTP, pH 7.35 with KOH, osmolarity 305 ± 5 mOsm/kg.

Whole-cell voltage-clamp recordings of transfected HEK293T cells were performed 48–72 hr after transfection using an EPC-10 amplifier and Patchmaster software (HEKA Elektronik, Lambrecht, Germany). Patch pipette electrodes with a tip resistance between 4 and 6 MΩ were pulled from thin-walled borosilicate glass capillaries (World Precision Instruments, Sarasota, FL) using a P-97 Flaming/Brown Puller (Sutter Instruments, Novato, CA). A reference Ag/AgCl wire was placed into the culture dish mounted on an inverted Olympus IX70 microscope. Recordings were performed at room temperature. A pressurized perfusion pencil multi-barrel manifold controlled with Valve Bank II (AutoMate Scientific, Inc, Berkeley, CA) was used to deliver extracellular solutions. ML218 of different concentrations (0.5, 1, 5, 25, and 100 mM) was added to the extracellular solution on the day of the experiments. Series resistance was compensated up to 70%, and passive membrane leak subtraction was conducted using a P/−4 protocol. Whole-cell Ca²⁺ currents (I_Ca) of transfected HEK293T cells were evoked for 50ms with incremental +5 mV steps from –80 mV to +65 mV. Current-Voltage (IV) data were fit with a single Boltzmann equation: I_Ca = G_max(V_m-V_r)/(1+exp[-(V_m-V_h)/k]), where G_max is the maximal conductance, V_m is the test voltage, V_r is the Ca²⁺ reversal potential, V_h is the membrane potential required to activate 50% of G_max, and k is the slope factor. Data were sampled at 100 kHz, filtered at 3 kHz, and analyzed using IgroPro (WaveMetrics).

Whole-cell voltage-clamp recordings of mouse cones, rods, and horizontal cells and macaque cone terminals were performed using an EPC-10 amplifier and Patchmaster software (HEKA). Patch pipette electrodes with a tip resistance between 10 and 14 MΩ for cones and 6–8 for horizontal cells were pulled from thick-walled borosilicate glass (1.5 mm outer diameter; 0.84 mm inner diameter; World Precision Instruments).

To prepare retinal slices, adult mice (6–8 weeks old) were anesthetized using isoflurane and euthanized by cervical dislocation. Eyes were enucleated, placed into cold Ames’ media slicing solution, and hemisected. Following the removal of the vitreous, the eye cup was separated into dorsal and ventral halves using a scalpel. Ventral retina was isolated, molded into low-melt agarose, and mounted in a Leica VT1200s vibratome (Leica Biosystems). Mouse retina slicing solution was continuously bubbled with 100% O₂ and contained the following: Ames’ Medium with L-glutamine supplemented with (in mM) 15 NaCl, 10 HEPES, 10 U/mL penicillin, 0.1 mg/mL streptomycin, pH 7.4, osmolarity 300 mOsm/kg. Vertical (~200 µm) or horizontal (~160 µm) retinal slices were anchored in a recording chamber, placed onto a fixed stage, and positioned under an upright Olympus BX51WI microscope equipped with a 60 X water-immersion objective (1.0 NA), and superfused with extracellular solution (flow rate of ~1–2 ml/min) at room temperature. Slices were visualized using IR-DIC optics and an IR-2000 (Dage MTI, Michigan City, IN) or SciCam Pro (Scientifica, Uckfield, United Kingdom) CCD camera controlled by the IR-capture software package or µManager, respectively (Edelstein et al., 2010; Edelstein et al., 2014). Drugs used in these experiments were added to the mouse extracellular solution on the day of experiments at the concentration described in Table 3. A reference Ag/AgCl pellet electrode was placed directly into the recording chamber solution. Data from whole-cell recordings with a series resistance >20 MΩ were discarded.

Cone and rod somas were identified based on their morphology and location. Cone identity was confirmed by the whole-cell capacitance (~3–4 pF), which is larger than rod whole-cell capacitance (~0.7–1 pF). Unless otherwise indicated, cones and rods were held at –90 mV for 200ms followed by a ramp of +0.5 mV/ms to +40 mV. To determine the voltage activation of I_Ca, whole-cell Ca²⁺ currents in cones were evoked for 50 ms with incremental +5 mV steps from –80 mV to +40 mV. The activation voltage of I_Ca is reported as G/G_max, where G is the conductance at each test voltage and G_max is the maximum peak conductance for each cone. Conductance was calculated using the equation I_Ca = G(V_m-V_r), where V_r is +60 mV. To determine steady-state inactivation of I_Ca, currents in cones were evoked for 500 ms with incremental +5 mV steps from –90 mV and –30 mV followed by a final step to –30 mV for 50 ms after each test voltage. The steady-state inactivation of I_Ca is reported as I/I_max, where I is the peak current in the final voltage step to –30 mV and I_max is the maximum peak current for each cone. Data were sampled between 20 and 60 kHz and filtered at 3 kHz.

For horizontal cell light responses, horizontal slices were prepared from the central mouse retina. The identity of horizontal cells was determined based on their larger soma diameter (~15 μm) compared to bipolar cell somas (~6 μm). During whole cell patch clamp recordings, horizontal cells were held at –70 mV. Light stimuli (1 s) at 410 nm (630 × 830 μm) were presented onto the retina (at a minimum of 5 s intervals) through the microscope’s condenser using a Polygon1000 DMD pattern illuminator (Mightex, Pleasanton, CA, USA) and a custom-built light path. Light intensity (in watts) was measured at the point on the microscope stage where the retina is placed using a power meter (Thorlabs, Newton, New Jersey, USA). Photon flux Ф_q (photons/s) within the light stimulus area was calculated using the measured light intensities in the formula:

where h is Planck’s constant (J*s), c is the speed of light (m/s), and λ is the wavelength (m). Light stimulus intensity was increased in Log2 steps from 4.9 × 10² to 2.1 × 10⁵ Ф_q/μm². For each light intensity step, both ON and OFF current amplitudes were measured from baseline (averaged 5 ms of current prior to light onset) to the maximum positive (after light onset) or maximum negative (after light offset) current, respectively. To confirm the identity of these horizontal cell light responses as AMPA-mediated currents, 1 s light stimuli (410 nm, 1.2 × 10⁵ Ф_q/μm²) were continuously delivered every 10 s before, during, and after bath application of 20 μM DNQX.

For voltage-clamp recordings of 13-lined ground squirrel cones, retinal slices were prepared as previously described (DeVries and Schwartz, 1999). The eyecup was divided along the dorsal to ventral axis into superior, middle, and inferior parts. The dorsal area above the line of the optic nerve head was defined as superior, the central region with a width of about 5 mm just ventral to the optic nerve head was middle, and the remaining ventral area was inferior. Isradipine and ML218, alone or in combination, were applied from separate puffer pipettes whose orifices were aimed at the cone synaptic region. Recordings were made with an Axopatch 200B amplifier (Molecular Devices). Signals were electronically filtered at 5 kHz and digitized at a rate of 10 kHz. Additional Gaussian filtering was added (cutoff frequency of 500 Hz). Tissue was viewed through a 63 x water immersion objective on a Zeiss Axioskop FS2 microscope and superfused with an extracellular solution at room temperature. Drugs and their concentrations used during these experiments are described in Table 3. Membrane potential was continuously maintained at –85 mV. During ramp stimulation, the membrane potential was depolarized to –85 to +35 mV at a rate of 1 mV/ms.

For rhesus macaque cone terminal recordings, a 12-year-old male was sedated with ketamine (5 mg/kg I.M.) and dexmedetomidine (0.015 mg/kg I.M.). Post-sedation, the animal received buprenorphine (0.02 mg/kg I.M.), atropine (0.02 mg/kg I.M.), and maropitant citrate (1 mg/kg S.Q.). The animal was intubated and maintained with inhaled isoflurane (0.75–2.0%) and propofol (7–8 mg/kg/hr I.V.). Crystalloid fluids (5 mL/kg/hr) were administered I.V., and phenylephrine (5–10 mcg/kg/hr I.V.) was used for blood pressure support. The animal was under anesthesia for approximately 3 hr prior to perfusion. Transcardial perfusion was approached through a midline thoracotomy. Immediately prior to perfusion, 5 mL of Euthasol (sodium pentobarbital 390 mg/mL/phenytoin sodium 50 mg/mL) was administered I.V. The descending thoracic aorta was clamped, the pericardium was opened, and the right atrium was cut. The apex of the left ventricle was sharply incised, and a large bore cannula (Yankauer suction handle, 5 mm internal diameter) was inserted through the left ventricle until it could be palpated in the ascending aorta. The cannula was clamped in place at the apex of the left ventricle. The animal was perfused with 4 L of cold phosphate-buffered saline at ~500 mL/min. Eyes were removed approximately 1 hr following perfusion. Eyes were dissected, and eye cups were allowed to dark adapt for ~30 min and stored in bicarbonate buffered Ames’ media at 32 °C equilibrated with 95% O₂/5% CO₂ prior to slice preparations. Vertical sections of central retina were prepared from 5 mm retina punches as described for mice. Depending on the species, the electrophysiological experiments were performed one or more times with cells from one animal representing technical replicates.

Retinas from 2 WT and 2 G369i KI mice (2–3 months old, all males) were dissected in ice-cold Dulbecco’s Phosphate Buffered Saline (D-PBS) and dissociated into single cells using the Papain Dissociation System (Worthington Biochemical). Two retinas were incubated in a 2.5 ml of dissociation solution (40 U/ml papain, 2 mM L-cysteine, 200 U/ml DNAse in Earle’s Balanced Salt Solution, EBSS) for 8 min at 37 °C. Following aspiration, 2 ml of inactivation solution (ovomucoid protease inhibitor and bovine serum albumin (BSA), 10 mg/ml each in EBSS) was added prior to trituration with a series of fire-polished, siliconized glass Pasteur pipettes. The solution was passed through a 40 μm mesh (pluriStrainer, Cat# 43-50040-51, pluriSelect) prior to centrifugation (200 × g for 5 min, 4 °C). Following aspiration of the supernatant, the cell pellet was resuspended in 0.5 ml D-PBS +0.5% BSA. Cell suspensions (10,000 cells per sample) were loaded on the Chromium Controller (10 X Genomics) and processed for cDNA library generation following the manufacturer’s instructions for the Chromium NextGEM Single Cell 3’ Reagent Kit v3.1 (10X Genomics). The resulting libraries were examined for size and quality using the Bioanalyzer High Sensitivity DNA Kit (Agilent) and their concentrations were measured using the KAPA SYBR Fast qPCR kit (Roche). Samples were sequenced on the NovaSeq 6000 instrument (paired end, read 1:28 cycles, read 2: 90 cycles) with a targeted depth of 9 K reads/cell.

scRNA-seq Fastq files were processed with CellRanger (v7.1.0) using the reference genome mm10–2020 A with introns included. CellRanger outputs were imported to Seurat (v5.0.1), a gene by feature count matrix was constructed, log normalization was performed, and the log normalized values were used for downstream analysis. Unsupervised clustering was performed in Seurat using resolution = 0.5. DEsingle method (v1.6.0, Miao et al., 2018) was used for differential expression analysis comparing G369i KI to WT in each cluster.

Eye cups were prepared from P42 WT and G369i KI littermates and fixed using 4% glutaraldehyde in 0.1 M cacodylate buffer, pH 7.4, for 4 hr at room temperature followed by additional fixation overnight at 4 °C. Glutaraldehyde-fixed eye cups were then washed 3 times in 0.1 M cacodylate buffer. Retinas were thereafter isolated and embedded in Durcupan resin after staining, dehydration, and embedding as described previously (Della Santina et al., 2016). A Thermo Scientific VolumeScope serial block face scanning electron micrscope was used to image embedded retinas. Retinal regions comprising a 2x2 montage of 40.96 μm tiles were imaged at a resolution of 5 nm/pixel and section thickness of 50 nm. Image stacks were aligned, and cone photoreceptor terminals were reconstructed using TrakEM2 (NIH). Postsynaptic partners at cone ribbons were followed to the inner nuclear layer to determine their identity. Amira software was used for 3D visualization of reconstructed profiles.

Retinal function was assessed using the Celeris system (Diagnosys, Inc) paired with the Espion software (Diagnosys, Inc). Mice were anaesthetized under red light (660  nm) via intraperitoneal (I.P.) injection with ketamine/xylazine mixture (100 mg/kg ketamine, 10 mg/kg xylazine). Tropicamide ophthalmic solution (1 %) and hypromellose lubricant eye gel (0.3%) were administered topically to both eyes before the mouse was secured to a heated (37 °C) platform to maintain body temperature. Ag/AgCl corneal stimulators were placed on each eye. After collecting data from individual mice, atipamezole (Antisedan, 1–2 mg/kg) was I.P. injection administered to reverse the effects of the ketamine and xylazine. For photopic ERGs, eyes were light adapted using a background green light at 20 cd·s/m² for 10 min. Following light adaptation, eight different light intensity pulses (–0.5, 0, 0.5, 1, 1.5, 2, 2.3, and 2.6 log cd·s/m²) were delivered on top of the background green light. ERG a-waves were measured from baseline to the peak of the negative potential. ERG b-waves were measured without filtering the oscillatory potentials from the peak of the a-wave to the peak of the positive potential within 60–70 ms (WT) and 80–100 ms (G369i KI and Ca_V1.4 KO) of the light stimuli. For flickering ERGs, mice were dark-adapted (>12 hr). White light pulses from –4–2.5 log cd·s/m² were delivered in 0.5 log unit steps at 10 Hz. Response amplitudes were measured from the trough to the peak of each response at all light intensities. Each intensity stimulus was delivered 10 times, with a 3 s interval between each stimulus, and averaged. The mean response amplitudes recorded in the right and left eye of each mouse are reported for all quantified ERG data. Experiments were performed at least three times with at least 3 mice per genotype (each mouse represented a biological replicate).

The visible platform swim test was performed as has been previously described (Bimonte-Nelson et al., 2015). WT, G369i KI, and Ca_v1.4 KO male and female mice (6–9 weeks old) were dark-adapted overnight and allowed to adapt to the procedure room for at least 1 hr prior to beginning the experiments. A water-filled, four-foot diameter galvanized steel tank and a visible white platform with a diameter of 10 cm was used for the swim test. Mice were subjected to six swim trials per day. Assays conducted under photopic (55 lux) and scotopic conditions (0 lux) were performed on days 1 and 2, respectively. Light intensity for photopic and scotopic conditions was measured at the platform using an Extech HD450 light meter (FLIR Systems, Nashua, New Hampshire). Mice were given 90 s to find the platform before being removed. After one trial was performed on all mice, the platform was moved to one of three different locations, top left, top center, and top right in relation to the initial site of mouse placement in the tank. Each trial was recorded using an infrared camera (Basler AG, Ahrensburg, Germany) and EthoVision XT16 software (Noldus Information Technology). Locating the platform was considered successful when mice contacted the platform with a head-on approach, even if mice failed to escape onto the platform. Mice were allowed to rest on the platform for 15 s at the end of each trial before being returned to a pre-warmed cage. Latency to find the platform was manually recorded and confirmed using recorded videos. Male and female mice were tested separately, and no sex differences in performance were identified post-hoc using two-way RM-ANOVA (Supplementary file 1). Male and female data were combined into a single group for each genotype. Latency to the platform was calculated by averaging the final 3 trials under scotopic and photopic conditions for each mouse and compared using Kruskal-Wallis one-way ANOVA with Dunn’s posthoc multiple comparison test. Experiments were performed at least three times with at least 3 mice per genotype (each mouse represented a biological replicate).

Retinas were dissected and stored in an oxygenated mouse retina-slicing solution. The loading of the cell-permeant Ca²⁺ dye Fluo-3, AM was modified from on a previous publication (Johnson et al., 2007). To label the base of cone terminals, retinas were incubated in retina slicing solution containing PNA Lectin conjugated with CF568 (100 $\mu g$/mL) for 15 min at room temperature. Then the retinas were incubated with retina slicing solution containing 1 mM Fluo-3, AM, and 0.025% pluronic acid (Biotium) for 45 min at room temperature followed by three rinses with retina slicing solution. The retinas were then incubated in retina slicing solution for 15 min at 35 °C to increase esterase activity. Vertical retina slices were prepared as described above. Slices were mounted on a Hyperscope multiphoton system (Scientifica, Uckfield, United Kingdom) equipped with a resonant scanner and a Mai Tai HP DeepSee laser (Spectra-Physics, Milpitas, CA, USA) tuned to 800 nm at 5% power. Slices were perfused with normal extracellular solution containing (in mM) 112 NaCl, 3 KCl, 1 MgCl₂, 3 CaCl₂, 1.25 NaH₂PO₄, 10 Na Pyruvate, 22.5 NaHCO₃, 5 HEPES, 10 Glucose, osmolarity 300 ± 5 mOsm/kg. High-K + extracellular solution contained (in mM) 65 NaCl, 50 KCl, 1 MgCl₂, 3 CaCl₂, 1.25 NaH₂PO₄, 10 Na Pyruvate, 22.5 NaHCO₃, 5 HEPES, 10 Glucose, osmolarity 300 ± 5 mOsm/kg. Extracellular solutions were supplemented with 20 μM CNQX, 25 μM L-APV, and 100 μM picrotoxin. Isradipine (10 μM) was added to High-K + extracellular solution. Images (512 × 512) were collected as 15-frame averages at 0.93 Hz using ScanImage (MBF Bioscience, Williston, VT, USA). Changes of Fluo-3 fluorescence in cone terminals were measured in ROIs drawn juxtaposed to PNA-CF568 labeling using Fiji (Schindelin et al., 2012). Experiments were performed at least three times with at least 3 mice per genotype (each mouse represented a biological replicate).

Electrophysiological data were analyzed by custom routines written in IgorPro software (Wavemetrics) and statistical analysis was performed using Prism software (GraphPad). Data were analyzed for normality by Shapiro Wilk test followed by parametric (t-test or ANOVA) or non-parametric methods (Kruskal Wallis, Mann-Whitney, Wilcoxon).

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

J W M, G J O, J D V, A H, K R, and A L were supported by NIH grants EY026817 (to A L), EY029953 (to J W M), and unrestricted funds from the University of Texas-Austin (to A L); S R W and M H were supported by NIH grant EY031677 (to M H), an unrestricted grant from RPB to UW Madison Dept. of Ophthalmology and a McPherson ERI professorship to M H; D F and S H D were supported by NIH grants EY012141 and EY032506, an unrestricted grant from RPB to the Northwestern University Dept. of Ophthalmology, and an RPB International Travel Award. N A S and R D Mayfield were supported by AA029955 (N A S) and AA0012404 and AA020926 (R D M). The authors thank J Sun for assistance with acquiring some of the electrophysiological data in Figure 3—figure supplement 1, S Knecht and R Wong for assistance with serial EM image collection, E Seidemann and B Shukla for the generous donation and collection of macaque retinal tissue, respectively, M Agosto for mGluR6 and TRPM1 antibodies, and M McCall and F Vinberg for advice on ERGs.

All animal experiments were performed in accordance with guidelines approved by the National Institutes of Health and the Institutional Animal Care and Use Committees at the University of Texas at Austin and Northwestern University were approved by the Institutional Animal Care and Use Committee (AUP-2023-00170).

1. Sent for peer review: December 5, 2023
2. Preprint posted: December 6, 2023
3. Reviewed Preprint version 1: February 2, 2024
4. Reviewed Preprint version 2: August 7, 2024
5. Reviewed Preprint version 3: August 28, 2024
6. Version of Record published: November 12, 2024

You can cite all versions using the DOI https://doi.org/10.7554/eLife.94908. This DOI represents all versions, and will always resolve to the latest one.

© 2024, Maddox, Ordemann et al.

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