ExSTED microscopy reveals contrasting functions of dopamine and somatostatin CSF-c neurons along the lamprey central canal

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Version of Record published: February 1, 2022 (This version)
Accepted: January 19, 2022
Received: August 17, 2021
Preprint posted: August 17, 2021 (Go to version)

1. Of interest
Cholinergic and noradrenergic axonal activity contains a behavioral-state signal that is coordinated across the dorsal cortex

Lindsay Collins, John Francis ... David A McCormick

Research Article Updated Jun 2, 2023
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Abstract

Cerebrospinal fluid-contacting (CSF-c) neurons line the central canal of the spinal cord and a subtype of CSF-c neurons expressing somatostatin, forms a homeostatic pH regulating system. Despite their importance, their intricate spatial organization is poorly understood. The function of another subtype of CSF-c neurons expressing dopamine is also investigated. Imaging methods with a high spatial resolution (5–10 nm) are used to resolve the synaptic and ciliary compartments of each individual cell in the spinal cord of the lamprey to elucidate their signalling pathways and to dissect the cellular organization. Here, light-sheet and expansion microscopy resolved the persistent ventral and lateral organization of dopamine- and somatostatin-expressing CSF-c neuronal subtypes. The density of somatostatin-containing dense-core vesicles, resolved by stimulated emission depletion microscopy, was shown to be markedly reduced upon each exposure to either alkaline or acidic pH and being part of a homeostatic response inhibiting movements. Their cilia symmetry was unravelled by stimulated emission depletion microscopy in expanded tissues as sensory with 9 + 0 microtubule duplets. The dopaminergic CSF-c neurons on the other hand have a motile cilium with the characteristic 9 + 2 duplets and are insensitive to pH changes. This novel experimental workflow elucidates the functional role of CSF-c neuron subtypes in situ paving the way for further spatial and functional cell-type classification.

Editor's evaluation

Here, the authors use a variety of optical super-resolution techniques to explore the structure and function of different neurons in tissue. They present very interesting evidence that sensory neurons contacting the cerebrospinal fluid in lamprey differ in the motility of their cilium and in their response to variations of pH: while somatostatin-positive ciliated sensory neurons lose dense core vesicles from their soma and enrich them in their axons, dopaminergic ciliated sensory neurons do not show any change in DSV localization / density. This manuscript is of broad interest for the neuroscience and imaging community.

https://doi.org/10.7554/eLife.73114.sa0

Introduction

The vertebrate spinal cord contains neurons with many different functions. In the middle, there is the central canal containing cerebrospinal fluid. The wall of the central canal is lined with ciliated cerebrospinal fluid-contacting (CSF-c) cells in all vertebrates (Agduhr, 1922; Vigh et al., 2004). In the lamprey, many CSF-c neurons are GABAergic sometimes with co-transmitters as somatostatin, neurotensin or dopamine (Christenson et al., 1991; Dale et al., 1997; Jalalvand et al., 2014; Rodicio et al., 2008). CSF-c neurons located at the lateral aspect of the central canal wall coexpress somatostatin and GABA and send axonal processes to mechanosensitive edge cells on the lateral margin of the spinal cord. GABA/somatostatin-expressing CSF-c neurons have recently been demonstrated to act as pH and mechanosensors and be part of a pH homeostatic system. At any deviation from neutral pH their activity is increased, which in turn leads to a depression of motor activity (Jalalvand et al., 2016a; Jalalvand et al., 2016b), which should counteract the processes that have led to the pH deviation. The accurate regulation of pH in the nervous system is of critical importance, and the fact that the GABA/somatostatin-expressing CSF-c neurons respond readily to pH changes gives them a particular homeostatic role. The effect of lowering the pH is blocked by APETx2, a specific acid-sensing ion channel 3 (ASIC3) antagonist (Diochot et al., 2004), as is their mechanosensitivity. The mechanism by which the GABA/somatostatin-expressing CSF-c neurons cause a reduction of locomotion has been suggested to be through release of somatostatin, since the effect on motor activity is blocked by somatostatin antagonists (Jalalvand et al., 2016a). CSF-c cells in a more ventral location contain dopamine (Brodin et al., 1990; Schotland et al., 1996) and their function is yet unknown.

In the present study, these two classes of CSF-c neurons, and their respective role for the operation of the pH homeostatic system and for the fluid transport in the central canal of the spinal cord is investigated. By applying a novel multi-scale imaging approach, using a high-throughput imaging method with sufficient resolution to resolve individual cells and organelles, we have been able to unravel the mode of operation of these two subtypes of CSF-c neurons. Light-sheet microscopy is the method of choice for rapid and minimally invasive acquisitions of large portions of tissues, but with a compromised spatial resolution in favour of recording speed and minimal photo-bleaching. To overcome this problem, we combine light-sheet with expansion microscopy (ExLSM) (Düring et al., 2019), a technique that physically expands tissue samples by a procedure including sample-embedding in a polyacrylamide gel and ends with ~four- to fivefold larger transparent samples (Chen, 2015; Tillberg et al., 2016). The physical expansion of the sample allows imaging of large volumes of spinal cord tissue with single-cell resolution and access to fine spatial information. This imaging approach allows quantification of the relative abundance and spatial patterns of CSF-c neuronal subtypes with specific focus on somatostatin- and dopamine-expressing cells along the central canal.

To further understand the physiological role of dopamine- and somatostatin-expressing CSF-c neurons, we take advantage of stimulated emission depletion (STED) microscopy to profile their neurotransmitter spatial distribution inside CSF-c neurons. STED microscopy (Hell and Wichmann, 1994; Willig et al., 2006) reaches a spatial resolution of ~40 nm, which allows us to resolve single synaptic vesicles even when the organelles are densely packed. This approach can identify neurotransmitter-specific release of dense-core synaptic vesicles during basal activity and upon pH stimulation in both somatostatin- and dopamine-expressing CSF-c neurons.

To gain structural information at ~5–10 nm level, an additional spatial resolution increase is needed, which we demonstrate with the application of STED imaging on expanded spinal cord tissues (ExSTED) (Gao et al., 2018). ExSTED imaging features an effective lateral spatial resolution of <10 nm, and allowed us to obtain cell-type-specific structural insight, previously accessible only with electron microscopy, on the cilia subtypes in ciliated somatostatin- and dopamine-expressing CSF-c. The cilia symmetry differs between primary (sensory) cilia (9 + 0 microtubule duplets) and motile cilia (9 + 2 microtubule duplets) and could be investigated in the context of specific cell types thanks to the spatial and specificity abilities of ExSTED. We could show that dopamine CSF-c neurons have motile cilia that may contribute to the flow of the cerebrospinal fluid, whereas somatostatin CSF-c neurons instead have predominantly sensory cilia conveying pH and mechanosensitivity.

Overall, this study uses recently developed high-resolution imaging techniques adapted to profiling the CSF-c neurons in the spinal cord. This provides cell-type information down to the molecular level. The experimental design developed in this study can be applied also to other types of tissue.

Results

Distribution of somatostatin- and dopamine-expressing CSF-c neurons along the spinal cord

To investigate the spatial distribution of CSF-c neurons around the central canal at the single-cell level with high speed and sufficient spatial resolution, we combined expansion and light-sheet microscopy on fluorescently labelled CSF-c neurons. An experimental design for expanding and handling spinal cord tissue was developed, which includes slicing, fluorescent immunolabelling of somatostatin and dopaminergic CFS-c neurons in 80–100 µm thick spinal cord slices, and the expansion steps with a final expansion factor of about ~4.5 (Figure 1A, B).

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Somatostatin and dopaminergic cerebrospinal fluid-contacting (CSF-c) neurons distribution along the spinal cord by expansion and light-sheet microscopy.

(A) A schematic illustration of the lamprey spinal cord treated for expansion microscopy (ExM). The spinal cords were immunostained for somatostatin and tyrosine hydroxylase (TH) prior to the ExM steps (MA-NHS treatment, gelation, proteinase K treatment, and expansion in water). (B) The spinal cord slices are shown before and after expansion. (C, D) Expanded samples imaged by light-sheet microscopy along the spinal cord. (C) Transverse and (D) horizontal images of somatostatin (magenta) and dopaminergic (green) CSF-c neurons shown by ExM-light-sheet microscopy. Scale bar, 30 µm. (E) Segmentation of the three-dimensional (3D) data from CSF-c neurons. Scale bar, 30 µm. (F) Quantification of somatostatin- and dopamine-expressing CSF-c neurons in four different areas of the spinal cord. The data are represented as the mean of number of cells in volume of each area; the error bar represents SD; Student’s paired t-test: *p ˂ 0.05 significant difference of somatostatin CSF-c neurons area 1 vs area 2 (p = 0.016, t₃ = −4.84), area 2 vs area 3 (p = 0.016, t₃ = 5.72) and vs area 4 (p = 0.04, t₃ = 3.38), **p ˂ 0.01 and ***p ˂ 0.001 significant difference of somatostatin and dopamine CSF-c neurons at area 1 (p = 5.8 × 10⁻³, t₃ = 7.06), at area 2 (p = 4 × 10⁻³, t₃ = 7.67), at area 4 (p = 7.9 × 10⁻⁴, t₃ = 13.9), and non-significant difference (n.s.) at area 3 (p = 0.09, t₃ = 2.40). cc, central canal.

Figure 1—source data 1 Distribution of somatostatin and dopaminergic cerebrospinal fluid-contacting (CSF-c) neurons along the spinal cord. Quantification of the somatostatin and dopaminergic CSF-c neurons in four areas (areas 1–4) of spinal cord.: https://cdn.elifesciences.org/articles/73114/elife-73114-fig1-data1-v1.xlsx
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The combination of our spinal cord expansion protocol and light-sheet microscopy (ExLSM) enables us to record a volume of 360 × 250 × 200 µm³ containing a large population of CSF-c neurons where the individual cells can be recognized and counted (Figure 1C, D). Somatostatin and dopaminergic (TH-expressing) CSF-c neurons can be visualized along the spinal cord (Figure 1C, D) and their specific location in the three-dimensional (3D) architecture of the tissue can be visualized (Figure 1—video 1) and quantified in different views (Figure 1E). The cell bodies of both dopamine and somatostatin CSF-c neurons are clearly visible, as well as their characteristic protrusion into the central canal. Different types of CSF-c neurons show a specific distribution along the lamprey spinal cord in all three spatial dimensions. Somatostatin-expressing CSF-c neurons are found throughout the whole volume and located laterally relative to the central canal, while the dopaminergic CSF-c neurons are located ventrally, confirming that the observation on individual sections is maintained in large volume with a high degree of order.

The throughput of ExLSM allowed recording a total volume of 7.2 × 10⁷ µm³ along the spinal cord for a total of 224 cells. We quantified the distribution of CSF-c neuronal subtypes at four different levels of the spinal cord, resulting in a higher amount of somatostatin CSF-c neurons compared to the dopaminergic CSF-c neurons in all four areas of the spinal cord. Additionally, in a specific position (area 2) the somatostatin CSF-c neurons were more abundant than in other parts (Figure 1F).

Somatostatin and dopamine neurotransmitters in CSF-c neurons are stored in dense-core vesicles

To investigate the subcellular location and compartmentalization of somatostatin and dopamine in CSF-c neurons, we used STED microscopy. The STED microscope was equipped with a glycerol objective and red-shifted wavelengths to allow tissue imaging with minimal spherical aberration and scattering.

Somatostatin and dopamine are stored in dense-core vesicles (DCVs) and were found in the soma, bulb protrusion (Figure 2A–C,E–G), and projections (data not shown) of the CSF-c neurons. No major differences between the somatostatin- and dopamine-positive puncta are observed. In both cases, the average diameter of somatostatin and dopamine DCVs was 100–120 nm with vesicles as small as 60 nm (full width at half maximum, FWHM; Figure 2D, H) measured with STED. However, using confocal microscopy, resulted in an overestimation of vesicle size with 250–300 nm (FWHM; Figure 2D, H). Our data on the somatostatin and dopamine DCVs diameter recorded with STED are compatible with previously published electron microscopy data (Schotland et al., 1996).

Figure 2

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Somatostatin and dopamine in cerebrospinal fluid-contacting (CSF-c) neurons are stored in dense-core vesicles.

(**A–C**) Somatostatin (magenta) and α-tubulin (green) immunostaining in CSF-c neurons. Scale bar in A, 5 µm. (**B, C**) Selected Region of interest (ROIs) of somatostatin dense-core vesicles (DCVs) in the bulb of somatostatin-expressing CSF-c neurons imaged with confocal and stimulated emission depletion (STED) microscopy, respectively. Scale bar, 500 nm. (D) Analysis of the size of somatostatin DCVs measured with confocal and STED microscopy (n = 46). (**E–G**) Dopamine (magenta) and phalloidin (green) immunostaining in CSF-c neurons. Scale bar in (E), 5 µm. (**F, G)** Selected ROIs of dopamine DCVs in the bulb of dopamine CSF-c neurons with confocal and STED microscopy, respectively. Scale bar, 500 nm. (H) Analysis of the size of dopamine DCVs with confocal and STED (n = 44). cc, central canal.

Figure 2—source data 1 Somatostatin in cerebrospinal fluid-contacting (CSF-c) neurons store in dense-core vesicles (DCVs). Analysis of somatostatin DCVs diameter measured with confocal and stimulated emission depletion (STED) microscopy.: https://cdn.elifesciences.org/articles/73114/elife-73114-fig2-data1-v1.xlsx
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Figure 2—source data 2 Dopamine in cerebrospinal fluid-contacting (CSF-c) neurons store in dense-core vesicles (DCVs). Analysis of dopamine DCVs diameter measured with confocal and stimulated emission depletion (STED) microscopy.: https://cdn.elifesciences.org/articles/73114/elife-73114-fig2-data2-v1.xlsx
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Somatostatin but not GABA release are responsible for pH response in somatostatin/GABA CSF-c neurons

The somatostatin CSF-c neurons in the lamprey spinal cord also express GABA (Brodin et al., 1990; Christenson et al., 1991; Jalalvand et al., 2014). We have recently shown that somatostatin/GABA CSF-c neurons are sensitive to pH changes of the cerebrospinal fluid (Jalalvand et al., 2016a; Jalalvand et al., 2016b). Here, we investigate how the spatial distribution and abundance of somatostatin and GABA changes during induced pH changes to acidic (6.5) or alkaline (8.5) conditions and if they were co-released. The somatostatin DCVs were visualized in the soma as well as in the axons by both confocal and STED microscopy (Figure 3A–F). Somatostatin DCVs were imaged with STED microscopy in control, acidic, and alkaline pH conditions. The number density of somatostatin DCVs measured for the soma of CSF-c neurons both in areas (Figure 3G) and volumes (Figure 3—figure supplement 1) decreased markedly in CSF-c neurons in either acidic or alkaline pH conditions. In contrast, the slices stained with a GABA antibody did not show any changes in fluorescence intensity at different pH (Figure 3H–K).

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Acidic and alkaline pH decreased the number of somatostatin dense-core vesicles (DCVs) in the soma but did not affect gamma-Aminobutyric acid (GABA) intensity.

(**A–F**) Spinal cord slices in normal (pH 7.4), acidic (pH 6.5), and alkaline (pH 8.5) extracellular solution stained with an anti-somatostatin antibody (magenta). (**A–C**) Confocal and stimulated emission depletion (STED) images (selected ROIs) of somatostatin DCVs in the soma. Scale bar in (A–C), 10 µm; in ROIs, 1 µm. (**D–F**) The axons of the somatostatin-expressing cerebrospinal fluid-contacting (CSF-c) neurons (arrowheads). Scale bar, 10 µm. (G) Quantification of somatostatin DCVs number density in cell area (µm⁻²) in the different conditions (n = 13). Student’s paired t-test: ***p ˂ 0.001 significant difference between pH 7.4 and 6.5 (p = 5.26 × 10⁻⁵, t₁₂ = 4.9), and 7.4 and 8.5 (p = 1.79 × 10⁻⁵, t₁₂ = 5.3). (**H–J**) The spinal cord slices in normal, acidic and alkaline extracellular solution, stained with an anti-GABA antibody (green). Scale bar, 10 µm. (K) Comparison of normalized GABA signals at pH 7.4 (n = 26), 6.5 (n = 24), and 8.5 (n = 22), respectively. Student’s t-test: non-significant difference (n.s.) between pH 7.4 and 6.5 (p = 0.62, t₄₇ = 0.48), and 7.4 and 8.5 (p = 0.80, t₄₃ = 0.25). (**L–O**) STED and confocal images of spinal cord slices stained for somatostatin (magenta) and GABA (green). (L) STED image of a CSF-c neuron. Scale bar, 1 µm. (**M, N**) Selected ROI from the soma in (l) shown at higher magnification with confocal and STED microscopy, respectively. Scale bar, 1 µm. (O) Line profile graph in image N. (P) Mean GABA signal in cellular compartments and compared to extracellular background (n = 5), normalized to volume intensity in soma (n = 5) and axons (n = 3), respectively. Repetitions are different cells. Student’s t-test between means of cellular means: ****p < 0.0001 significant difference between soma-volume and background (p = 1.0 × 10⁻⁵, t₈ = −9.7), soma-DCVs and background (p = 1.0 × 10⁻⁵, t₈ = −9.6), axon-volume and background (p = 8.0 × 10⁻⁸, t₄ = −93), and axon-DCVs and background (p = 4.4 × 10⁻⁵, t₄ = −19), non-significant differences (n.s.) between soma-volume and DCVs (p = 0.090, t₈ = −1.9), and axon-volume and DCVs (p = 0.12, t₄ = 1.9). Data (**G, K, P**) are represented as means, with error bars representing standard deviation (SD) (**G, K**) or standard error of the mean (SEM) (P). cc, central canal.

Figure 3—source data 1 Effect of acidic or alkaline pH on somatostatin dense-core vesicles (DCVs) number density in cell area of somatostatin-expressing cerebrospinal fluid-contacting (CSF-c) neurons. Quantification of somatostatin DCVs number density in cell area of somatostatin-expressing CSF-c neurons (µm^–2) in the different pH.: https://cdn.elifesciences.org/articles/73114/elife-73114-fig3-data1-v1.xlsx
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Figure 3—source data 2 Effect of acidic or alkaline pH on GABA signal in somatostatin-expressing cerebrospinal fluid-contacting (CSF-c) neurons. Analysis of the GABA intensity signal in somatostatin-expressing CSF-c neurons in the different pH.: https://cdn.elifesciences.org/articles/73114/elife-73114-fig3-data2-v1.xlsx
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Figure 3—source data 3 No correlation between GABA and somatostatin signals in somatostatin-expressing cerebrospinal fluid-contacting (CSF-c) neurons. Analysis of GABA signal in and outside of somatostatin dense-core vesicles (DCVs) in soma and axon of somatostatin-expressing CSF-c neurons.: https://cdn.elifesciences.org/articles/73114/elife-73114-fig3-data3-v1.xlsx
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As somatostatin and GABA are co-expressed in the same CSF-c neurons, we investigated if they were colocalized in the same vesicles (Figure 3L–P). STED images of somatostatin DCVs and GABA-expressing CSF-c neurons did not show colocalization of somatostatin DCVs and GABA in the soma (Figure 3M–O). The GABA signal was not significantly different when measured in or outside of somatostatin DCVs, neither in the soma nor in the axons (Figure 3P). The results confirmed that there is no correlation between GABA and somatostatin signals in somatostatin vesicles. Our imaging data support the conclusion that during pH changes of the extracellular solution vesicles containing somatostatin are released, resulting in fewer DCVs in the soma, while GABA is not co-released.

Dopaminergic CSF-c neurons are not sensitive to changes in extracellular pH

The next step was to explore whether the dopaminergic CSF-c neurons at the ventral aspect of the central canal were also sensitive to pH changes. As for the somatostatin experiments, acidic and alkaline extracellular pH was perfused on spinal cord slices and subsequently stained with a dopamine antibody to investigate the dopamine DCVs spatial distribution in dopaminergic CSF-c neurons. We performed STED microscopy to resolve and count the single vesicles in distinct neuronal locations (Figure 4A–C). The result showed no significant change in the number density of dopaminergic DCVs after perfusion with acidic or alkaline pH solutions, neither in cell areas (Figure 4D) nor in cell volumes (Figure 4—figure supplement 1).

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Dopaminergic cerebrospinal fluid-contacting (CSF-c) neurons did not respond to acidic and alkaline pH.

(**A–C**) Stimulated emission depletion (STED) images of dopamine-containing dense-core vesicles (DCVs) in the soma of CSF-c neurons in normal (pH 7.4), acidic (pH 6.5), and alkaline (pH 8.5) extracellular solution. Scale bar, 1 µm. (D) Quantification of the number of dopamine DCVs number density in cell area (µm⁻²) in the different conditions (n = 10). Student’s t-test: non-significant (n.s.) between pH 7.4 and 6.5 (p = 0.27, t₉ = 1.12), and 7.4 and 8.5 (p = 0.29, t₉ = 1.08). (E) Whole-cell patch recording of a CSF-c neuron, showing firing spontaneous action potentials in control (pH 7.4), acidic (p H 6.5), and alkaline (pH 8.5) conditions in the presence of gabazine (20 mM) and kynurenic acid (2 mM). (**F–H**) Photomicrographs of the CSF-c neurons recorded in (E) intracellularly filled with Neurobiotin (arrow) during recording. The labelled cell showed immunoreactivity to tyrosine hydroxylase (TH, arrow). Scale bar, 10 µm. (I) Action potential frequency during 1 min in CSF-c neurons at pH 7.4, 6.5, and 6.8, respectively (n = 15). Student’s paired t-test: non-significant difference (n.s.) between pH 7.4 and 6.5 (p = 0.24, t₁₄ = −1.22), and 7.4 and 8.5 (p = 0.1, t₁₄ = −1.75). The bar graph data are represented as the means, with error bars representing standard deviation (SD). cc, central canal.

Figure 4—source data 1 Effect of acidic or alkaline pH on dopamine dense-core vesicles (DCVs) number density in cell area of dopaminergic cerebrospinal fluid-contacting (CSF-c) neurons. Quantification of dopamine DCVs number density in cell area (µm⁻²) in dopaminergic CSF-c neurons in the different pH.: https://cdn.elifesciences.org/articles/73114/elife-73114-fig4-data1-v1.xlsx
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Figure 4—source data 2 Effect of acidic and alkaline pH on action potential frequency in dopaminergic cerebrospinal fluid-contacting (CSF-c) neurons. Quantification of action potential frequency in dopaminergic CSF-c neurons at different pH conditions during 1 min.: https://cdn.elifesciences.org/articles/73114/elife-73114-fig4-data2-v1.xlsx
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To complement the results from the STED imaging, dopaminergic CSF-c neurons were patched as previously described for somatostatin-expressing CSF-c neurons (Jalalvand et al., 2016a; Jalalvand et al., 2016b). Their electrophysiological properties and response to changes of extracellular pH were investigated with whole-cell patch recording in current-clamp mode (Figure 4E). All recorded dopaminergic CSF-c neurons fired spontaneous action potentials, depolarizing synaptic potentials, and hyperpolarizing synaptic potentials (Figure 4E). To verify that responses were not evoked synaptically, the GABAergic and glutamatergic synaptic transmission was blocked by bath-application of gabazine (GABA_A receptor antagonist) and kynurenic acid (glutamate receptor antagonist). During bath-applied extracellular solutions of acidic pH (6.5) or alkaline pH (8.5) no changes in spike frequency were observed, nor a net depolarization of the resting membrane potential (Figure 4E1). Thus, in contrast to the somatostatin CSF-c neurons the dopamine CSF-c are not sensitive to pH changes. All neurons labelled with Neurobiotin during the patch-clamp recording were tyrosine hydroxylase (TH) immunoreactive (Figure 4F, H).

Both somatostatin and dopamine CSF-c neurons are ciliated (see below) and the former known to be mechanosensitive (Jalalvand et al., 2016a). To test if the dopaminergic CSF-c neurons are mechanosensitive very brief fluid pulses were applied near their bulb protrusion in the central canal as previously done for the somatostatin CSF-c neurons (Figure 4—figure supplement 2A; Jalalvand et al., 2016b). Dopamine CSF-c neurons responded with an action potential and a distinct mechanosensitive response (Figure 4—figure supplement 2B). To investigate if the mechanosensitivity in dopaminergic CSF-c neuron was mediated by the ASIC3 as in somatostatin CSF-c neurons, APETx2, a specific blocker of ASIC3, was applied. In the presence of APETx2 we still observed a mechanosensitive response (Figure 4—figure supplement 2C). Thus, the mechanosensitivity is not mediated by ASIC3 in dopamine CSF-c neurons. In situ hybridization showed expression of polycystic kidney disease 2-like 1 (PKD2L1) channels in dopaminergic CSF-c neurons (Figure 4—figure supplement 2D). As PKD2L1 has been confirmed as a mechanosensitive ion channel in zebrafish (Böhm et al., 2016; Sternberg et al., 2018), the results suggest that the mechanosensitivity of the dopaminergic CSF-c neurons may be mediated by PKD2L1 channels. In contrast, the mechanical response of somatostatin CSF-c neurons is blocked by an ASIC3 antagonist.

In conclusion, dopamine CSF-c neurons are not sensing pH changes as their somatostatin counterparts, but both are mechanosensitive. The mechanical transduction is blocked by an ASIC3 antagonist in somatostatin CSF-c neurons and may be mediated by PKD2L1 in the dopamine CSF-c neurons as in the zebrafish.

CSF-c neurons show both primary and motile cilia symmetries

Somatostatin- and dopamine-expressing CSF-c neurons are thus both mechanosensitive, but through different transduction mechanisms. Since both types of CSF-c neurons are ciliated, we investigated whether the functional difference is reflected in the structural organization of the cilia.

Cilia express acetylated α-tubulin and can be classified either as primary cilia, mainly present in sensory cells and neurons and with a 9 + 0 symmetry of nine outer microtubule doublets, or motile cilia, with a 9 + 2 symmetry showing an extra pair of microtubule singlets in the centre (Gaertig and Wloga, 2008; Satir, 2005). Primary and motile cilia both have an average diameter of 200–240 nm, close to the achievable spatial resolution of a confocal microscope equipped with a high numerical aperture objective. Therefore, a higher spatial resolution is crucial to assess their organization and subcompartmentalization.

One aim for exploring the cilia symmetry of ciliated CSF-c neurons was to uncover potential signalling compartments related to pH and mechanosensitivity of the CSF-c neurons. Cilia in the central canal was immunolabelled for acetylated α-tubulin, a protein characteristic of cilia, and imaged with confocal and STED microscopy to measure their diameters, respectively (Figure 5A, B, E, F). STED imaging visualizes the cilium as a hollow structure with an outer diameter of about 240 nm thanks to the increased spatial resolution, but the resolution is still not enough to detect the microtubule doublets. One strategy to increase the observable level of detail is to expand the tissue. In the expanded spinal cord tissue, pre-immunostained for acetylated α-tubulin, the resolution of the dense cilia in the central canal increased (Figure 5—figure supplement 1, 3D XYZ image) as compared to confocal imaging of the non-expanded sample. Confocal imaging of the expanded sample shows spatial details comparable to STED images (Figure 5C) of the non-expanded one, therefore still not having the resolution (5–10 nm) needed to dissect the internal cilia structure and identify microtubule doublets. That level of detail is crucial to further investigate the nature of cilia as sensory or motile for each specific cell type.

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Primary and motile cilia symmetry are present in the lamprey spinal cord.

(**A, B**) Confocal and stimulated emission depletion (STED) images of a cerebrospinal fluid-contacting (CSF-c) neuron cilium in a non-expanded spinal cord pre-stained with anti-α-tubulin antibodies. Scale bar, 0.5 µm. (**C, D**) Confocal (expansion microscopy, ExM) and STED (ExSTED) images of a cilium in the expanded spinal cord. Scale bar, 0.5 µm. (**E, F**) Quantification of the cilium diameter (arrowheads) in confocal and STED images in a non-expanded spinal cord. (G, H) Quantification of the cilium diameter (arrowheads) in confocal (ExM) and STED (ExSTED) in the expanded spinal cord. (**I–M**) Confocal (ExM, **I, J**) and STED (ExSTED, K, L) deconvoluted images of a primary cilium with 9 + 0 symmetry in the expanded spinal cord. Scale bar, I, 2 µm and J–L, 1 µm. (M) Quantification of the cilium diameter from image (l) (arrowheads). (**N–R**) Confocal (ExM, **N, O**) and STED (ExSTED, P, Q) deconvoluted images of a motile cilium with 9 + 2 symmetry in the expanded spinal cord. Scale bar, N, 2 µm and O–Q, 1 µm. (R) Quantification of the cilium diameter from image (Q) (arrowheads). Cilia diameters in blue were divided by the expansion factor.

Figure 5—source data 1 Quantification of a cilium diameter of cerebrospinal fluid-contacting (CSF-c) neuron with confocal microscopy in non-expanded spinal cord.: https://cdn.elifesciences.org/articles/73114/elife-73114-fig5-data1-v1.xlsx
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Figure 5—source data 2 Quantification of a cilium diameter of cerebrospinal fluid-contacting (CSF-c) neuron with stimulated emission depletion (STED) microscopy in non-expanded spinal cord.: https://cdn.elifesciences.org/articles/73114/elife-73114-fig5-data2-v1.xlsx
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Figure 5—source data 3 Quantification of a cilium diameter of cerebrospinal fluid-contacting (CSF-c) neuron with confocal microscopy in expanded spinal cord (expansion microscopy, ExM).: https://cdn.elifesciences.org/articles/73114/elife-73114-fig5-data3-v1.xlsx
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Figure 5—source data 4 Quantification of a cilia diameter of cerebrospinal fluid-contacting (CSF-c) neuron with stimulated emission depletion (STED) microscopy in expanded spinal cord (ExSTED).: https://cdn.elifesciences.org/articles/73114/elife-73114-fig5-data4-v1.xlsx
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Figure 5—source data 5 Quantification of a primary cilium diameter of cerebrospinal fluid-contacting (CSF-c) neuron with 9 + 0 symmetry with stimulated emission depletion (STED) microscopy in expanded spinal cord (ExSTED).: https://cdn.elifesciences.org/articles/73114/elife-73114-fig5-data5-v1.xlsx
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Figure 5—source data 6 Quantification of a motile cilium diameter of cerebrospinal fluid-contacting (CSF-c) neuron with 9 + 2 symmetry with stimulated emission depletion (STED) microscopy in expanded spinal cord (ExSTED).: https://cdn.elifesciences.org/articles/73114/elife-73114-fig5-data6-v1.xlsx
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By using the combination of STED microscopy and expanded tissue (ExSTED), we were able to add a factor of ~four to five to the typical 50 nm resolution of STED microscopy and therefore resolving at the smaller spatial scale of 5–10 nm (Figure 5—figure supplement 2). In this way, it was possible to separate high-density cilia within the 3D geometry of the central canal (Figure 5—video 1) due to the increased imaging resolution and further resolve their internal structure and allowing to classify them as motile or sensory (Figure 5D). In some cilia, the central pair of tubules was clearly observed with a peak-to-peak distance of 70 nm (Figure 5P–R).

Both types of cilia symmetries in the lamprey spinal cord were explored: sensory (Figure 5I–M) and motile cilia symmetries (Figure 5N–R). The presence of motile cilia in the central canal of the lamprey has been shown with electron microscopy (Schotland et al., 1996; Vigh et al., 2004), but with ExSTED we could detect both motile and sensory cilia symmetries in this area. As all CSF-c neurons, including somatostatin and dopaminergic CSF-c neurons in the lamprey spinal cord, are ciliated, the next step is to know which cilia type is related to which neuronal phenotypes and if they contribute to a specific neuronal function.

Somatostatin CSF-c neurons have both primary and motile cilia, but dopaminergic CSF-c neurons have only motile cilia

To further investigate whether somatostatin-expressing CSF-c neurons (pH-sensitive) and dopaminergic CSF-c neurons (non-pH-sensitive) have different types of cilia, expansion was combined with STED microscopy. Using dual-colour imaging of spinal cord slices, either somatostatin or dopamine (TH immunostaining), together with α-tubulin immunostaining (Figure 6) was analysed. To have better access to the cilia and CSF-c neurons the central canal of the expanded spinal cord has been cut horizontally and flipped 90° with respect to the coverslip. The central canal can then be visualized in a cylindrical form (Figure 6A–C). To confirm what type of CSF-c neurons the cilia belong to, z-stack scanning for all the imaging was applied. In our data, 70 cilia belong to somatostatin-expressing CSF-c neurons (n = 70). Of these, 60 cilia were primary (9 + 0 symmetry) and 10 cilia were motile (9 + 2 symmetry) (Figure 6D–K, T). Additionally, 20 cilia of dopaminergic CSF-c neurons were all motile cilia (9 + 2 symmetry) (Figure 6L–S, T). We found that somatostatin-expressing CSF-c neurons are sensory neurons expressing mainly sensory cilia. However, the motile cilia in both somatostatin-expressing and dopaminergic CSF-c neurons might be involved in contributing to the flow of CSF through the central canal (Figure 6U).

Figure 6 with 1 supplement see all

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Cilia symmetries in somatostatin and dopaminergic cerebrospinal fluid-contacting (CSF-c) neurons.

(A) A schematic illustration of an expanded spinal cord stained for somatostatin (magenta), dopamine (blue), and α-tubulin (green). The gel was cut through the central canal and flipped 90° on the side on the coverslip. (**B, C**) Longitudinal images of the expanded spinal cord (expansion microscopy, ExM) stained with α-tubulin, TH, and somatostatin antibodies, respectively. Scale bar, 10 µm. (**D–K**) ExM and ExSTED images of two somatostatin CSF-c neurons with their cilia. Scale bar, D, 1 µm and H, 3 µm. (**F, G**) showing 9 + 0 symmetry and (J, K) showing both 9 + 0 and 9 + 2 symmetries. Scale bar, 1 µm. (**L–S**) ExM and ExSTED images of two dopaminergic CSF-c neurons (TH staining) with 9 + 2 symmetry. Scale bar, L–O, Q–S, 1 µm and P, 2 µm. (T) Quantification of cilium types in somatostatin and dopaminergic CSF-c neurons. (U) A schematic illustration of the central canal with somatostatin and dopaminergic CSF-c neurons and their possible cilia symmetries. cc, central canal.

The sensory cilia are a potential location for pH-sensitive receptors (Atkinson et al., 2019; Vina et al., 2015). We have recently shown that in somatostatin-expressing CSF-c neurons, acidic and alkaline responses are mediated by the ASIC3 and possibly PKD2L1 channel, respectively (Jalalvand et al., 2016a; Jalalvand et al., 2016b). Interestingly, we could detect the expression of both ASIC3 and PKD2L1 on the cilia of spinal CSF-c neurons in mice (Figure 6—figure supplement 1A–Q). Besides, we visualized Arl13b, a ciliary protein on cilia that has high expression in sensory cilia (Figure 6—figure supplement 1R–V).

CSF neurons have one and rarely two cilia

Another advantage of expansion microscopy (ExM) is that more specific morphological details of cells can be revealed, which cannot be detected in non-expanded samples. Using ExM, paired cilia of CSF-c neurons were visualized in some cases (Figure 7). As reported, somatostatin-expressing and dopaminergic CSF-c neurons mainly have one cilium, but we could in rare cases (N = 8) detect two cilia per bulb of these neurons by ExM (Figure 7A–F). To confirm the finding, we performed z-stack scanning to see the whole bulb and its cilia in 3D (Figure 7—video 1).

Figure 7 with 1 supplement see all

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Cerebrospinal fluid-contacting (CSF-c) neurons have one or two cilia on their bulb protrusions.

(A) An expansion microscopy (ExM) image of an expanded somatostatin CSF-c neuron showing one cilium on its bulb (arrow). Scale bar, 5 µm. (B, C) A somatostatin CSF-c neuron with two cilia (arrow) (C) selected ROI from (B). Scale bar B, 5 µm and C, 2 µm. (D) A dopaminergic CSF-c neuron (tyrosine hydroxylase [TH] expressing) with one cilium (arrow). Scale bar, 5 µm. (E, F) A dopaminergic CSF-c neuron with two cilia (arrows) (F) selected ROI from (E). Scale bar, E, 5 µm and F, 2 µm. cc, central canal.

Discussion

In this study, we uncovered the dense spatial organization of somatostatin- and dopamine-expressing CSF-c neurons along the central canal. ExLSM allowed us to visualize and quantify CSF-c neurons in a large volume of the spinal cord with a fourfold increase in resolution, necessary to distinguish and count each individual cell. There were approximately twice as many somatostatin CSF-c neurons as dopaminergic CSF-c neurons along the whole length of the spinal cord. One area, at the level of the rostral part of the dorsal fin had, however, a somewhat larger number of somatostatin CSF-c neurons than elsewhere. This is the same spinal cord area in which a dramatic decrease occurs of the serotonin/dopamine neurons that are located just ventral to the central canal (Figure 8A, Schotland et al., 1995; Zhang et al., 1996). Whether this is a coincidence or a related phenomenon remains to be elucidated.

Figure 8

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Somatostatin and dopaminergic cerebrospinal fluid-contacting (CSF-c) neurons are two distinct cell types with contrasting function along the spinal cord.

(A) Schematic illustration of a cross-section of the lamprey spinal cord, with a somatostatin- and dopamine-expressing CSF-c neuron at the central canal and their axonal projections. (B) Summary of phenotypes, organelles, and physiological properties of dopaminergic and somatostatin CSF-c neurons.

Somatostatin and dopamine DCVs were quantified with STED microscopy (Figure 8B). The size distribution is in line with previous electron microscopy studies (Schotland et al., 1996). The STED data reveal the presence of DCVs in several CSF-c cellular subcompartments, including the bulb protrusion into the central canal, soma, and axonal branches. We could show a marked reduction of somatostatin DCVs number density in the soma of CSF-c neurons when exposed to acidic or alkaline pH, while there was no effect on the dopamine DCVs number density. This lack of change in the number density of DCVs in dopaminergic CSF-c neurons agrees with the present electrophysiological results showing that these cells are not activated by changes of the extracellular pH.

We have shown previously that somatostatin/GABA-expressing CSF-c neurons are responding to acidic and alkaline extracellular solutions (Jalalvand et al., 2016a; Jalalvand et al., 2016b) being part of a homeostatic system for reducing motor activity when the spinal cord/brain is exposed to changes of pH as due to metabolic or respiratory stress or intense muscle activity. We now show with STED imaging that the pH response resulted from somatostatin DCVs release, whereas GABA was unaffected.

At the ultrastructural level, GABAergic CSF-c neurons have been shown to have ciliated endings in the lamprey and other species (Orts-Del’Immagine et al., 2014; Schotland et al., 1996; Vigh et al., 2004). Since somatostatin/GABA and dopaminergic/GABA CSF-c neurons have similar responses to mechanical stimuli, while only the former responds to pH changes, the question arose if these two types of CSF-c neurons have different forms of cilia. To visualize cilia microtubule symmetry in CSF-c neurons a resolution of 5–10 nm is needed, which was provided by the combination of expansion and STED microscopy (ExSTED) (Gao et al., 2018). ExSTED microscopy allowed us to obtain cell-type-specific structural insights on the cilium types in ciliated somatostatin- and dopamine-expressing CSF-c neurons, respectively (Figure 8B). Using ExSTED microscopy we found primary cilia with 9 + 0 symmetry in somatostatin-expressing CSF-c neurons, also found in specialized sensory cells (Kirschen and Xiong, 2017; Singla and Reiter, 2006) and motile cilia with 9 + 2 symmetry in dopaminergic CSF-c neurons that may serve to generate fluid flow (Faubel et al., 2016; Satir and Christensen, 2007). The high expression of sensory cilia (9 + 0) found in pH/mechanosensitive somatostatin-expressing CSF-c neurons provide additional evidence to recognize them as sensory neurons. The mechanosensitivity of the dopamine- and somatostatin-expressing CSF-c neurons is mediated by different cellular mechanisms, the latter is blocked by APETx2, a selective ASIC3 blocker as shown in previous studies (Jalalvand et al., 2016a; Jalalvand et al., 2016b). The mechanosensitivity of the dopamine CSF-c neurons is, however, unaffected by APETx2. This blocker is specific to the ASIC3 but does not block ASIC1 receptors (Diochot et al., 2004), the latter having been cloned in lamprey (Coric et al., 2005) and found insensitive to pH. The data support the previous interpretation that ASIC3 mediates both the acid-sensing and the mechanosensitivity, in somatostatin-expressing CSF-c neurons (Jalalvand et al., 2016a; Jalalvand et al., 2016b). PKD2L1 channel is expressed in the dopaminergic CSF-c neurons (Figure 4—figure supplement 2) and this channel is known to serve as a mechanosensitive receptor (Kamura et al., 2011; Nauli et al., 2003) and may therefore serve in this function also for the dopaminergic CSF-c neurons. In the zebrafish, the mechanosensitive GABA CSF-c neurons sense the lateral bending movements occurring during locomotion through PKD2L1 channels (Fidelin et al., 2015). We have also observed expression of ASIC3 and PKD2L1 channels, as well as sensory cilia in CSF-c neurons of the mouse, which suggests that this may be representative of all vertebrates. The motile cilia (9 + 2) found only in the mechanosensitive (non-pH) dopaminergic CSF-c neurons may suggest that they may act as a CSF flow generator.

In conclusion, by applying ExM combined with light-sheet and STED microscopy with nanoscale precision, the spatial organization, abundance, and subcellular composition of two distinct GABAergic CSF-c neuronal subtypes in the lamprey spinal cord were elucidated. One type is sensitive to pH and displaying mechanosensitivity (somatostatin) while the other (dopamine) only responds to mechanical stimuli. They use different cellular mechanisms for the transduction of mechanical stimuli to their cilia.

Materials and methods

Key resources table

Reagent type (species) or resource	Designation	Source or reference	Identifiers	Additional information
Biological sample (Lampetra fluviatilis)	Spinal cord	Collected from the Ljusnan River, Hälsingland, Sweden		Freshly isolated from Lampetra fluviatilis
Biological sample (Mus musculus)	Spinal cord	Janvier Labs, C57BL/6		Freshly isolated from C57BL/6
Antibody	Anti-acetylated tubulin (mouse monoclonal)	Sigma-Aldrich	Cat# T6793, RRID:AB_477585	IF (1:500)
Antibody	Anti-somatostatin (rat monoclonal)	Millipore	MAB354, RRID:AB_2255365	IF (1:100)
Antibody	Anti-somatostatin 14-IgG (rabbit polyclonal)	Peninsula laboratories	Cat# T-4102.0400, RRID:AB_518613	IF (1:1000)
Antibody	Anti-TH (mouse monoclonal)	Millipore	Cat# MAB318, RRID:AB_2201528	IF (1:200)
Antibody	Anti-TH (rabbit polyclonal)	Millipore	Cat# AB152, RRID:AB_390204	IF (1:500)
Antibody	Anti-dopamine (mouse monoclonal)	Millipore	Cat# MAB5300, RRID:AB_94817	IF (1:400)
Antibody	Anti-GABA (mouse monoclonal)	Swant	Cat# Mab 3A12, RRID:AB_2314454	IF (1:2000)
Antibody	Anti-polycystin-L (rabbit polyclonal)	Millipore	Cat# AB9084, RRID:AB_571091	IF (1:500)
Antibody	Anti-ASIC3 (rabbit polyclonal)	Thermo Fisher Scientific	Cat# PA5-41022, RRID:AB_2576713	IF (1:400)
Antibody	Anti-ARL13B (rabbit polyclonal)	Proteintech	Cat# 17711-1-AP, RRID:AB_2060867	IF (1:500)
Antibody	Donkey-anti-rat-IgG-AF594	Jackson ImmunoResearch Labs	Cat# 712-585-153, RRID:AB_2340689	IF (1:500)
Antibody	Donkey-anti-rat-IgG-AF488	Jackson ImmunoResearch Labs	Cat# 712-545-153, RRID:AB_2340684	IF (1:200)
Antibody	Goat-anti-mouse-STAR635P	Abberrior	Cat# ST635P-1001-500 UG, RRID:AB_2893232	IF (1:500)
Antibody	Goat-anti-rabbit-AF594	Thermo Fisher Scientific	Cat# A-11037, RRID:AB_2534095	IF (1:500)
Antibody	Donkey-anti-mouse-IgG-Cy3	Jackson ImmunoResearch Labs	Cat# 715-165-150, RRID:AB_2340813	IF (1:500)
Antibody	Donkey-anti-mouse-IgG-AF488	Jackson ImmunoResearch Labs	Cat# 715-545-150, RRID:AB_2340846	IF (1:200)
Other	NeuroTrace530/615	Thermo Fisher Scientific	Cat# N21482, RRID:AB_2620170	IF (1:1000)
Other	NeuroTrace640/660	Thermo Fisher Scientific	Cat# N21483, RRID:AB_2572212	IF (1:1000)
Other	Phalloidin-STAR635P	Abberior		IF (1:200)
Peptide, recombinant protein	Neurobiotin	Vector Laboratories	Cat# SP-1120, RRID:AB_2313575	Injection of 0.5% solution for intracellular labelling
Peptide, recombinant protein	Streptavidin-AF488	Jackson ImmunoResearch	Cat# 016-540-084, RRID:AB_2337249	IF (1:1000)
Chemical compound, drug	Glutamate receptor antagonist kynurenic acid	Tocris Ellisville, MO, USA		Bath perfusion, 2 mM
Chemical compound, drug	GABAA receptor antagonist gabazine	Tocris Ellisville, MO, USA		Bath perfusion, 20 mM
Software, algorithm	Fiji	Schindelin et al., 2012	RRID:SCR_002285
Software, algorithm	MATLAB	The Mathworks	RRID:SCR_001622
Software, algorithm	Impsector	Max-Planck Innovation	RRID:SCR_015249
Software, algorithm	Origin	OriginLab	RRID:SCR_014212
Software, algorithm	Imaris 9.1	Bitplane	RRID:SCR_007370
Software, algorithm	Clampex and Clampfit	Molecular Devices, CA, USA	RRID:SCR_011323
Commercial assay or kit	Digoxigenin RNA Labeling kit	Roche Diagnostics	Catalog #11 277 073 910	In situ hybridization
Commercial assay or kit	TSA Cy3 Plus Evaluation Kit	PerkinElmer	NEL763E001	In situ hybridization
Antibody	Anti-DIG antibody coupled to HRP (sheep polyclonal)	Roche Diagnostics	RRID: AB_514497	IF (1:2000)

Animals: lamprey

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Experiments were performed on a total of 40 adult river lampreys (Lampetra fluviatilis) of both sexes that were collected from the Ljusnan River, Hälsingland, Sweden. The experimental procedures were approved by the local ethical committee (Stockholms Djurförsöksetiska Nämnd; Dnr 5806-2019) and were in accordance with The Guide for the Care and Use of Laboratory Animals (National Institutes of Health, 1996 revision). During the investigation, every effort was made to minimize animal suffering and to reduce the number of animals used during the study.

Mouse

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Experiments were performed on a total of 4, C57BL/6 wild type mice (Mus musculus). All experiments were performed in accordance with animal welfare guidelines set forth by Karolinska Institutet and were approved by Swedish Board of agriculture for Animal welfare (ethical permit number: 2645-2021).

Immunohistochemistry: lamprey

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The animals (n = 30) were deeply anesthetized through immersion in carbonate-buffered tap water containing MS-222 (100 mg/l; Sigma, St Louis, MO, USA). Following decapitation, a portion of the spinal cord, rostral to the dorsal fin, was fixed by 4% (wt/vol) paraformaldehyde (PFA) in phosphate-buffered saline (PBS) for 12–24 hr at 4°C, and subsequently cryoprotected in 20% sucrose in phosphate buffer (PB) for 3–12 hr. For GABA and dopamine immunodetection, 1% glutaraldehyde (vol/vol) was added to the fixative solution. Transverse sections (20 µm thick) were cut on a cryostat (Microm HM 560, Microm International GmbH, Walldorf, Germany), collected on gelatine-coated slides, and kept at −20°C until processing. Sections were incubated/co-incubated with different primary antibodies listed here: a mouse monoclonal anti-acetylated tubulin antibody (dilution 1:500, Sigma-Aldrich), a rat monoclonal anti-somatostatin antibody (1:100, Millipore), a rabbit polyclonal anti-somatostatin-14 IgG antibody (1:1000, Peninsula laboratories), a mouse monoclonal anti-TH antibody (1:200, Millipore), a rabbit polyclonal anti-TH antibody (1:500, Millipore), a mouse monoclonal anti-dopamine antibody (1:400, Millipore), and a mouse monoclonal anti-GABA antibody (1:2000, Swant).

Mouse

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The animals were deeply anesthetized with sodium pentobarbital (200 mg/kg i.p.) and transcardially perfused with 4% PFA in 0.01 M PBS, pH 7.4. The spinal cord was removed and postfixed for 2 hr, after which it was transferred to a 12% sucrose solution in 0.01 M PBS overnight. Transverse sections (20 µm thick) were cut on a cryostat and mounted on gelatine-coated slides and kept at −20°C until processing. Sections were incubated with a rabbit polyclonal anti-polycystin-L antibody (1:500, Millipore), a rabbit polyclonal ASIC3 antibody (1:400, Thermo Fisher Scientific), or a rabbit polyclonal anti-ARL13B antibody (1:500, Proteintech).

The lamprey and mouse spinal cord sections were after incubation with primary antibodies thoroughly rinsed in PBS and then incubated with Alexa Fluor 594-conjugated donkey anti-rat IgG (1:500, Jackson ImmunoResearch Labs) or Alexa Fluor 488-conjugated donkey anti-rat IgG (1:200, Jackson ImmunoResearch Labs), STAR 635P-conjugated goat anti-mouse (1:500, Abberior) or Alexa Fluor 594-conjugated donkey anti-mouse (1:500, Thermo Fisher Scientific), and STAR 635P-conjugated goat anti-rabbit (1:500, Abberior) or Alexa Fluor 594-conjugated goat anti-rabbit (1:500, Thermo Fisher Scientific), for 3 hr at room temperature. The sections were Nissl stained by adding NeuroTrace 530/615 red or 640/660 deep-red fluorescent Nissl stain (1:1000, Invitrogen) to the secondary antibody solution. Phalloidin conjugated with STAR 635P (1:200, Abberior) was added to the secondary antibodies to stain actin filaments. The primary and secondary antibodies were diluted in 1% bovine serum albumin and 0.3% Triton X-100 in 0.1 M PB. All sections were mounted in custom-made Mowiol mounting media, supplemented with DABCO (Thomas Scientific, C966M75), and covered with coverslips (No. 1.5).

Expansion microscopy

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After fixation, transverse sections of the lamprey spinal cord (40–50 µm thick) were cut on a cryostat and collected and immersed in PBS. The sections were stained with antibodies according to the protocols described above. The samples were treated at room temperature for 1 hr with anchoring solution, 1 mM MA-NHS in PBS (0.018 g MA-NHS in 100 µl DMSO and stored at −20°C) which enables proteins to be anchored to the hydrogel. The slices then were incubated for 1 hr in a monomer solution on ice, which was followed by adding the gelling solution to the gelling chambers. The gelling solution consisted of monomer solution (1 M NaCl, 8.6% sodium acrylate, 2.5% acrylamide, and 0.15% N,N′-methylene bisacrylamide in PBS), 4-hydroxy-TEMPO (0.2%), Tetramethylethylenediamine (TEMED) (accelerator solution, 0.2%), and Ammonium persulfate (APS) (initiator solution, 0.2%). The slices were incubated in a 37°C incubator for 2 hr for gelation. The gels were removed from the gelling chamber and the coverslips were transferred to digesting buffer (50 mM Tris, pH 8.0, 1 mM EDTA, and 0.5 Triton X-100) with proteinase K (1:100, 8 units/ml, New England Biolabs) for 30–40 min in room temperature. The gels were removed from the digestion buffer and immersed in deionized (DI) water (three to five times for 30 min) for further expansion. After final expansion, the gels were mounted to a 35 mm glass bottom petri dish coated with poly-L-lysine (Sigma-Aldrich). To remove the extra gel on a coverslip and increase the resolution, we cut the gel through the central canal and rotated it 90° away from the coverslip. The expansion factor was ~4.5–5 and has been calculated by overlapping the pre- and post-expanded gel slice in the air–water boundary and by cross checking the cilia diameter in expanded and not samples.

STED microscopy

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Most STED images have been recorded on a custom-built setup, as previously described (Alvelid and Testa, 2019) equipped with a glycerol objective (HCX PL APO 93×/1.30 NA GLYC STED White motCORR, 15506417, Leica Microsystems). The images were recorded by exciting Alexa Fluor 594 and STAR 635P with 561 nm (PDL561, Abberior Instruments) and 640 nm (LDH-D-C-640, PicoQuant) laser lines, respectively. A STED beam at 775 nm (KATANA 08 HP, OneFive) has been used to deplete both laser lines, shaped by a spatial light modulator (LCOS-SLM X10468-02, Hamamatsu Photonics) into a donut. Two-colour STED images were recorded line-by-line sequentially. A third confocal channel, for structures labelled with Alexa Fluor 488, has been excited with a 510-nm laser line (LDH-D-C-510, PicoQuant). Detection was performed using the following bandpass filters: ET615/30 m (Chroma), ET705/100 m (Chroma), and FF01-550/49 (Semrock). The pixel sizes of the images were 27.2–32.3 nm in XY, and 300 nm in Z. The pixel dwell time used was 10–50 µs, added over one to two lines. Laser powers used were in the following ranges: 561 nm: 1–40 µW, 640 nm: 2–20 µW, and 775 nm: 65–180 mW. Additional STED images (Figures 2 and 3A–J) have been recorded on a commercial Leica TCS SP8 3X STED. The images were recorded by exciting Alexa Fluor 488, Alexa Fluor 594, and STAR 635P with laser lines at 488, 594, and 635 nm, respectively. The detection windows used were 510–560, 600–645, and 670–750 nm. The pixel size of the images was 25–30 nm, and the pixel dwell time used was 10–40 µs, added over two to four lines.

Light-sheet microscopy

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The expanded slices from four different parts of the lamprey spinal cord were separately glued to a metal rod and placed in the chamber of a Zeiss Light-sheet Z1 microscope containing DI water. Fluorescence was excited from two sides using ×10/0.2 NA illumination objectives and detected using a ×10/0.4 NA or ×20/1.0 NA water dipping objective.

Image analysis

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The images were analysed using Fiji (Schindelin et al., 2012), Imspector (Max-Planck Innovation), and MATLAB (The Mathworks). The transverse sections of the cilia images were deconvoluted with a calculated effective STED PSF, Lorentzian with FWHM of 50 nm, using the Richardson–Lucy deconvolution implemented in Imspector. The deconvolution was stopped after five iterations. The line profiles of cilia and DCVs were fitted with a Gaussian, and the FWHM was measured. The OriginLab software (OriginLab) was used for making the graphs. Animation and spot object creation tools of Imaris 9.1 (Bitplane) were used to make 3D movies and segmentation of the 3D data.

Analysis of the correlation of GABA signal and somatostatin DCVs (Figure 3v) was performed by manually selecting areas inside (soma/axon-DCVs) and outside (soma/axon-volume) somatostatin DCVs (detected in the somatostatin channel) as well as in the extracellular space (Background). The mean of the GABA signal inside each area was recorded. A total of five cells were used for the soma and three cells for the axons. The number of areas selected in each category in each cell ranged from 3 to 63, with a mean of 37. Plotted are the paired graphs for each cell, with each data point representing the cellular mean of the mean GABA signals per area. The Student’s t-test is performed between the groups of means of the cells per category.

The quantification of the somatostatin and dopamine DCVs in the cell area (2D) was performed using Fiji Analyze particles plugin. Vesicles with an area smaller than 0.190 µm² (corresponding to vesicle diameter less than 250 nm) were considered and their number density was calculated.

The quantification of the somatostatin and dopamine DCVs in cell volume was performed using the 3D imageJ suite plugins (Ollion et al., 2013). The 3D stacks were slightly adjusted for noise applying a two-pixel 3D median filter. The background was detected by applying a 15-pixel 3D median filter and then subtracted from the stacks. The stacks were then segmented by first detecting the seeds points using the maximum local filter and then applying the 3D spot segmentation. Vesicles with a volume smaller than 0.008 µm³ were considered for the final quantification and their number density was calculated.

Live slices

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The spinal cord of lamprey (n = 5) was dissected out and embedded in 4% agar dissolved in ice-cooled oxygenated 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES)-buffered physiological solution containing (in mM): 138 NaCl, 2.1 KCl, 1.8 CaCl₂, 1.2 MgCl₂, 4 glucose, 2 HEPES, and with pH adjusted to 7.4 with NaOH. The agar block containing the spinal cord was glued to a metal plate and transverse slices of the spinal cord (100–150 or 300 µm) were cut on a vibrating microtome. The preparation was continuously perfused with HEPES solution at 4–6°C. Then the spinal cord slice was exposed to HEPES solution with various pH values (7.4, 6.5, and 8.5) for 8–10 min. The slices were then fixed immediately with 4% PFA (somatostatin) or 4% PFA and 1% glutaraldehyde (dopamine and GABA) in 0.01 M PBS overnight at 4°C. Following a thorough rinse in 0.01 M PBS, the slices (100–150 µm) were incubated with a rat monoclonal anti-somatostatin, a mouse monoclonal anti-GABA, or a mouse monoclonal anti-dopamine antibody overnight at 4°C. The slices were then incubated with Alexa Fluor 594-conjugated donkey anti-rat or anti-mouse IgG as described above.

Patch-clamp recordings

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The spinal cord slices (300 µm) of lamprey (n = 5) after slicing were transferred in a cooled recording chamber and allowed to recover at 5°C for 1 hr before recording. Patch electrode was prepared from borosilicate glass microcapillaries (Hilgenberg GmbH) using a two-stage puller (PP830, Narishige, Japan). Patch electrodes (8–12 MΩ) were filled with an intracellular solution of the following composition (in mM): 130 K-gluconate, 5 KCl, 10 HEPES, 4 Mg-ATP, 0.3 Na-GTP, and 10 phosphocreatine sodium salt. The pH of the solution was adjusted to 7.4 with KOH and osmolarity to 270 mOs ml⁻¹ with water. Cells ventral to the central canal were recorded in whole-cell in current-clamp mode using a Multiclamp 700B amplifier (Molecular Devices Corp., CA, USA). Bridge balance and pipette capacitance compensation were adjusted and signals were digitized and recorded using Clampex software and analysed in Clampfit (pCLAMP 10, Molecular Devices, CA, USA). The neurons were visualized with DIC/infrared optics (Olympus BX51WI, Tokyo, Japan). Resting membrane potentials were determined in current-clamp mode during whole-cell recording. The following drugs were added to the extracellular solution and applied by bath perfusion: the GABAA receptor antagonist gabazine (20 mM, Tocris, Ellisville, MO, USA) and the glutamate receptor antagonist kynurenic acid (2 mM, Tocris, Ellisville, MO, USA). Neurons were intracellularly labelled by injection of 0.5% Neurobiotin (Vector Laboratories) during whole-cell recordings. After recording the spinal cord slices were fixed with 4% formalin. To investigate whether the intracellularly Neurobiotin-labelled CSF-c cells express TH, the slices were incubated overnight with a mouse monoclonal anti-TH antibody and then rinsed thoroughly in 0.01 M PBS and incubated with a mixture of Alexa Fluor 488-conjugated streptavidin (1:1000, Jackson ImmunoResearch Labs) and Cy3-conjugated donkey anti-mouse IgG (1:500, Jackson ImmunoResearch Labs) for 3 hr at room temperature.

In situ hybridization

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Lamprey (n = 2) were deeply anesthetized as described above and the rostral spinal cord was removed, fixed in 4% formalin in 0.1 M PB overnight at 4°C, and then cryoprotected in 20% sucrose in 0.1 M PB. Then, 10–20 µm thick cryostat sections were made and stored at −80°C until processed. Single-stranded digoxigenin-labelled sense and antisense PKD2L1 riboprobes were generated by in vitro transcription of the previously cloned PKD2L1 cDNA using the Digoxigenin RNA Labeling kit (Roche Diagnostics). Briefly, sections were incubated for 1 hr in prehybridization mix (50% formamide, 5× SSC, 1% Denhardt’s, 50 g/ml, salmon sperm DNA, 250 g/ml yeast RNA) at 60°C. Sections incubated with the heat-denatured digoxigenin-labelled riboprobe were hybridized overnight at 60°C. Following the hybridization, the sections were rinsed twice in 1× SSC, washed twice in 1× SSC (30 min each) at 60°C and twice in 0.2× SSC at room temperature. After blocking in 0.5% blocking reagent (PerkinElmer), the sections were incubated overnight in anti-DIG antibody coupled to HRP (1:2000, Roche Diagnostics) at 4°C. The probe was then visualized by TSA Cy3 Plus Evaluation Kit (PerkinElmer). The specificity of the hybridization procedure was verified by incubating sections with the sense riboprobe (data not shown). The sections were rinsed thoroughly in 0.01 M PBS and then incubated with a mouse monoclonal anti-TH antibody (1:200, Millipore) overnight at 4°C, rinsed in PBS, and incubated with Alexa Fluor 488-conjugated donkey anti-mouse IgG (1:200, Jackson ImmunoResearch Labs) and 640/660 deep-red fluorescent Nissl stain (1:1000, Thermo Fisher Scientific) for 2 hr and mounted with Mowiol. All primary and secondary antibodies were diluted in 1% BSA and 0.3% Triton X-100 in 0.1 M PB.

Data availability

Data availability The data that support the implementation of the method and support the findings in this study, including images and statistical analysis are openly available in Zenodo with reference number 5758671 at https://doi.org/10.5281/zenodo.5758671.

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Decision letter

Suzanne R Pfeffer

Senior and Reviewing Editor; Stanford University School of Medicine, United States
Francesca Bottanelli

Reviewer
Claire Wyart

Reviewer; Institut du Cerveau et la Moelle épinière, Hôpital Pitié-Salpêtrière, Sorbonne Universités, UPMC Univ Paris 06, Inserm, CNRS, France

Our editorial process produces two outputs: (i) public reviews designed to be posted alongside the preprint for the benefit of readers; (ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Decision letter after peer review:

Thank you for submitting your article "ExSTED microscopy reveals contrasting functions of dopamine and somatostatin CSF-c neurons along the central canal" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, and the evaluation has been overseen by Suzanne Pfeffer as the Senior and Reviewing Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Francesca Bottanelli (Reviewer #1); Claire Wyart (Reviewer #2).

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential revisions:

1. The introduction should give more insights into the underlying biology and also clarify which questions are being asked here: what is new and why these are important, to help the reader understand the biology that underlies this work.

2. The interpretation of physiological results based on established knowledge of ASICs in lamprey cannot be easily explained, and the authors need to provide further data to support their claims or rephrase their conclusions with great care. Specific details follow in the individual reviews below. Electrophysiological recordings of both SST+ and DA+ neurons is important to make conclusions related to the functional roles of these cells, as the microscopy alone cannot infer on the sensory function; alternatively the text needs to be modified in line with suitable conclusions from the morphological analyses.

3. Many of the figures were provided in suboptimal resolution and it is essential to be sure they are provided at the appropriate quality.

Reviewer #1 (Recommendations for the authors):

The work is generally well presented. However, some additional information could make it easier for the non-expert reader. Here are some suggestions:

• A model at the end to compile all results on pH sensitivity, mechanosensing, etc… would greatly simplify the understanding of the messages of the paper.

• TH staining is not explained.

• The authors claim a resolution below 10 nm however the size of the antibodies used for staining should be considered.

• The authors say "Somatostatin-expressing CSF-c neurons are found throughout the whole volume and located laterally relative to the central canal, while the dopaminergic CSF-c neurons are located ventrally" could the author point that out in figure 1 as I have a hard time visualizing it.

• Figure 1f = Cell number per volume? Or total cell number?

• Figure 3g = vesicles per volume? Or total amount of vesicles per soma? Additionally, the increase of number of vesicles in axon is not quantified?

• Figure 4d = same comment as for figure 3.

• Figure 5 = I am wondering why the expansion factor was calculated by measuring the gel slice rather than looking at a sub-cellular structure. Could the authors calculate the expansion factor by measuring cilia diameter in STED and expanded confocal samples?

Reviewer #2 (Recommendations for the authors):

1. On the microscopy section, the authors make a statement that DSVs within somatostatin-positive sensory cells are present in the axons at baseline and become more numerous upon pH variations. To illustrate their point, the authors need to show:

a) Images illustrating that DSVs were found in the axonal extension (page 5, "data not shown") at baseline, and;

b) Evidence determining whether DSVs are released or transported from the soma to the axon (Figure 3d-f) upon changes of pH. A quantification of the total number of DSV per cell could be insightful to resolve these 2 interpretations.

This point is important for interpretation of the mechanisms at play upon variations of pH.

a) the authors need to show that somatostatin-positive cells respond to pH in their conditions as it is key for their interpretation, and their previous article on all spinal CSF-contacting neurons did not mention that only half the ciliated neurons contacting the CSF responded to pH (Jalalvand et al. Current Biology 2016).

b) Similarly, the authors here examine the mechanosensory response of dopaminergic CSF-contacting neurons without quantifying the response of somatostatin-positive neurons to the same stimulus. Both responses to variations of pH and mechanical stimulation need to be performed in both cell types.

c) Another group have shown that lamprey only express one type of ASIC channel, ASIC1 (not the channel ASIC3 ): see https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3047259/). Therefore, how can the authors explain the effect of the drug used against ASIC3 in this context ?

In order to solve this issue, the authors could show expression of ASIC channels, and test the effect of a drug against ASIC1 on the response of somatostatin-expressing cells.

d) Finally, the only ASIC channel expressed in lamprey, ASIC1, when cloned and tested, has been reported to be proton insensitive (see https://www.ncbi.nlm.nih.gov/pmc/articles/PMC1464184/)--how then to explain then the activation of a subset of ciliated neurons exposed to protons?

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "ExSTED microscopy reveals contrasting functions of dopamine and somatostatin CSF-c neurons along the lamprey central canal" for further consideration by eLife. Your revised article has been reviewed by 3 peer reviewers and the evaluation has been overseen by Suzanne Pfeffer as the Senior and Reviewing Editor.

We would like to publish this story in eLife but ask you to please make textual changes in response to this reviewer's comments.

Specific comments from one reviewer:

The authors satisfactorily answered some of the issues raised on the quantification of images and provided high resolution images for their outstanding data acquired at high resolution. These data are very convincing and beautiful. Congratulations, this will be very precious and inspirational for further studies in the spinal field !

The issue remaining are related to the interpretation of the physiological results. The authors should either show expression of ASIC3 in CSF-cNs by ISH or tone down their conclusions relative to the ASIC3 receptor in response to pH.

Similarly the authors should be more careful for stating the role PKD2L1 in pH sensing or mechanoreception as they lack genetic methods to investigate its function in lamprey.

A) On the role of ASIC3 in pH sensing :

As stated in my first review, ASIC3 can only be found in the genome of mammals. Only ASIC1 has been identified in the lamprey genome (https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3047259) and the lamprey ASIC1a is proton insensitive (https://www.ncbi.nlm.nih.gov/pmc/articles/PMC1464184/).

The authors did not respond to my request of testing the role of ASIC1a in their experiments using pharmacology.

One can therefore wonder whether 2uM APETx2 used by the authors (about 100X the EC50 of ASIC3 homomers, https://pubmed.ncbi.nlm.nih.gov/17113616/) could be acting on other targets (see suggested actions on Na or K channels mentioned here, https://pubmed.ncbi.nlm.nih.gov/17113616/). Please discussion clearly in the text.

B) On the role of PKD2L1 in mechanoreception:

The authors summarize that PKD2L1 is responsible for mechanoreception only in dopaminergic CSF-cNs.

However:

1. PKD2L1 is expressed in all CSF-cNs, ventral (dopaminergic) and dorsal (somatostatinergic) as shown by the authors here and found in zebrafish (Djenoune 2014) and mouse (Huang 2006; Petracca 2018). In fact, in their 2016b publication, the authors had proposed that SST+ CSF-cNs were responding to basic pH via PKD2L1, suggesting a role for this channel in these cells.

Why therefore in this study, only mentioning the role of PKD2L1 for mechanoreception in the dopaminergic CSF-cNs despite the expression being there in both dorsal and ventral cell types?

2. Due to the lack of specific antagonists, the authors do not have the tools in lamprey to measure its contribution to mechanoreception. They can only suggest that PKD2L1 contribute to mechanoreception in both ventral and dorsal CSF-cN types.

3. Please correct an error in citation and references used:

The authors use Bohm et al. 2016 to state that CSF-cNs rely on PKD2L1 to be mechanosensory in zebrafish.

To be correct, we showed in vivo in Bohm 2016 that both ventral and dorsal CSF-cNs respond to concave (not convex) mechanical deformations of the spinal cord via PKD2L1 (response are abolished in the KO).

However, mechanoreception cannot be rigorously demonstrated in vivo. It is therefore only in Sternberg 2018 that we could show in vitro using a piezo device to mechanically stimulate their membrane that all CSF-cNs isolated in primary cultures are mechanosensory cells and that their response always rely on PKD2L1.

Note that in zebrafish, we now understand that CSF-cNs in vitro do not respond to CSF flow (Prendergast et al. under review) and in vivo, their response to concave mechanical bending of the spinal cord needs their interaction with the Reissner fiber (Orts Dell Immagine 2020), which does not alter itself the flow (Cantaut-Belarif 2018).

The authors should at minima cite Sternberg et al. 2018 for showing the role of PKD2L1 in mechanoreception, but to be fair, also propose with more nuances in the discussion how these cells in lamprey can combine ASIC and PKD channels to sense pH and mechanical inputs, citing Bohm 2016, Sternberg 2018 and Orts Del Immagine 2020.

https://doi.org/10.7554/eLife.73114.sa1

Author response

Essential revisions:

1. The introduction should give more insights into the underlying biology and also clarify which questions are being asked here: what is new and why these are important, to help the reader understand the biology that underlies this work.

We have rewritten part of the Introduction to better expose the underlying biological questions.

2. The interpretation of physiological results based on established knowledge of ASICs in lamprey cannot be easily explained, and the authors need to provide further data to support their claims or rephrase their conclusions with great care. Specific details follow in the individual reviews below. Electrophysiological recordings of both SST+ and DA+ neurons is important to make conclusions related to the functional roles of these cells, as the microscopy alone cannot infer on the sensory function; alternatively the text needs to be modified in line with suitable conclusions from the morphological analyses.

We have previously reported on the sensitivity of the somatostatin-CSF-contacting neurons (SST+) specifically showing that they respond to both fluid movements and pH changes by electrophysiology (Jalalvand et al. 2016, Nature Communications, Jalalvand et al. 2016, current Biology). We now performed similar experiment on dopaminergic CSF-c neurons in the current study. Since the responses of SST+CSF neurons have been quantified in considerable detail earlier, we feel that there would be no reason to repeat these results in the context of these experiments.

3. Many of the figures were provided in suboptimal resolution and it is essential to be sure they are provided at the appropriate quality.

We provide Figures at high resolution.

Reviewer #1 (Recommendations for the authors):

The work is generally well presented. However, some additional information could make it easier for the non-expert reader. Here are some suggestions:

• A model at the end to compile all results on pH sensitivity, mechanosensing, etc… would greatly simplify the understanding of the messages of the paper.

A new figure, Figure 8, has been added to summarise our findings.

• TH staining is not explained.

Dopaminergic CSF-c neurons express Tyrosine hydroxylase (TH). TH catalyzes the hydroxylation of the Tyrosine to L-DOPA (precursor of Dopamine). Anti-TH antibody has been used to detect TH and is as a marker of dopaminergic neurons. In the Method/Immunohistochemistry section we have explained the type of used TH antibodies and the method of the staining.

• The authors claim a resolution below 10 nm however the size of the antibodies used for staining should be considered.

The 10 nm value was derived by the response of our STED system to point-like objects, such as single antibodies, which are imaged with a FWHM of ~35 10 nm, and further divided by the expansion factor of 4. However, this value refers to how good the response is, which is only one metric to define spatial resolution. When considering biological structures, we agree that the size of the labels indeed matters and influences the resulting ability of resolving and visualizing fine structures. In this work, to be able to tell apart the duplets, we needed at least a resolution of 10-15 nm, which was achieved with ExSTED but not with only STED.

• The authors say "Somatostatin-expressing CSF-c neurons are found throughout the whole volume and located laterally relative to the central canal, while the dopaminergic CSF-c neurons are located ventrally" could the author point that out in figure 1 as I have a hard time visualizing it.

We added in Figure 1 the annotation for dorsal and ventral to guide the viewer to the geometry of the central canal.

• Figure 1f = Cell number per volume? Or total cell number?

We have counted somatostatin and dopamine CSF-c cells in volume of 1.8 × 10⁷ µm³ in each part of the spinal cord. What we see in the graph shows the mean number of the respected cells per volume of each part. We have counted a total of 224 cells in 4 distinct parts of the spinal cord within a volume of 7.2 × 10⁷ µm³. (Information added in figure 1 legend)

• Figure 3g = vesicles per volume? Or total amount of vesicles per soma? Additionally, the increase of number of vesicles in axon is not quantified?

We have clarified the quantification presented in figure 3g in the figure legend. The somatostatin DCVs number density (µm^-2) is reported per cell soma in N = 13 cells. These are 2DSTED images with a pixel size of 20-25 nm and an axial resolution of ~500 nm. (Information added in the Figure legend). We have additionally applied volumetric imaging with 2D STED and quantified somatostatin and dopamine DCVs densities as the number N within the soma volume (N x µm^-3) (added in Figure 3—figure supplement 1 and Figure 4—figure supplement 1). Quantification of the density of vesicles in axons have not been included since we found it hard to have a comprehensive view of the entire axonal network.

• Figure 4d = same comment as for figure 3.

As above we quantified vesicle density in soma areas (N x µm^-2) (2DSTED), and quantification vesicle number density of dopamine DCVs in volume (N x µm^-3) has been added in Figure 4—figure supplement 1 (Information added in the Figure legend).

• Figure 5 = I am wondering why the expansion factor was calculated by measuring the gel slice rather than looking at a sub-cellular structure. Could the authors calculate the expansion factor by measuring cilia diameter in STED and expanded confocal samples?

We agree that a direct check in the sample structure is more sensitive to local change in expansion as well as isotropicity compared to the overall gel especially for the study that precise amount of expansion factor is important. However, to measure the diameter of the same cilia before and after expansion is almost impossible regarding the numerous and packed cilia in the spinal cord samples. In addition to calculating the expansion factor by measuring the size of the sample/gel before and after expansion, we measured the diameter of several cilia (before and after expansion) and calculated the expansion factor by using the means of the diameter from STED and ExSTED. We also made a graph for comparison of cilia diameters (divided by the expansion factor in the expanded one) in the different techniques (the graph added in the Figure 5—figure supplement 2).

Reviewer #2 (Recommendations for the authors):

1. On the microscopy section, the authors make a statement that DSVs within somatostatin-positive sensory cells are present in the axons at baseline and become more numerous upon pH variations. To illustrate their point, the authors need to show :

a) Images illustrating that DSVs were found in the axonal extension (page 5, "data not shown") at baseline, and;

Not visible in Figure 2 but in Figure 3 D-F.

b) Evidence determining whether DSVs are released or transported from the soma to the axon (Figure 3d-f) upon changes of pH. A quantification of the total number of DSV per cell could be insightful to resolve these 2 interpretations.

This point is important for interpretation of the mechanisms at play upon variations of pH.

We quantified the number density of vesicles per soma area (µm^-2) (2DSTED) for somatostatin and dopamine DCVs (explained above). We have additionally applied volumetric imaging and quantified the number density of somatostatin and dopamine DCVs within soma volumes (µm^-3) added in Figure 3—figure supplement 1 and figure 4—figure supplement 1.

2. The interpretation of physiological results based on established knowledge of ASICs in lamprey cannot be easily explained, and the authors need to provide further data to support their claims or rephrase their conclusions:

a) the authors need to show that somatostatin-positive cells respond to pH in their conditions as it is key for their interpretation, and their previous article on all spinal CSF-contacting neurons did not mention that only half the ciliated neurons contacting the CSF responded to pH (Jalalvand et al. Current Biology 2016).

The study of Jalalvand et al. 2016 dealt with one specific subtype of CSF-contacting cells that express somatostatin and are located at the lateral aspect of the central canal and has axonal projections to the lateral margin of the spinal cord (see also new Figure 8). It was known from previous studies that another subtype of CSF-contacting neurons in lamprey expressed dopamine (Schotland et al., 1996), in which the cells have a more ventral location. The latter with a previously unknown function has now been included in the present study and a function unravelled.

b) Similarly, the authors here examine the mechanosensory response of dopaminergic CSF-contacting neurons without quantifying the response of somatostatin-positive neurons to the same stimulus. Both responses to variations of pH and mechanical stimulation need to be performed in both cell types.

We have previously reported on the sensitivity of the somatostatin-CSF-contacting neurons (Jalalvand et al. 2016, Nature Communications) specifically showing that they respond to both fluid movements and pH changes, and that both responses are blocked by APETx2, a specific ASIC3 antagonist. Since these responses have been quantified in considerable detail earlier, we feel that there would be no reason to repeat these results in the context of these experiments.

c) Another group have shown that lamprey only express one type of ASIC channel, ASIC1 (not the channel ASIC3 ): see https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3047259/). Therefore, how can the authors explain the effect of the drug used against ASIC3 in this context ?

We note that in the article of Coric et al. 2005, they had identified one clone of cDNA that corresponded to ASIC1 and when expressed in oocytes, no pH sensitivity was found. They do not comment regarding the possible presence of ASIC3. The review of Grunder and Chen (2010) has a focus on ASIC1a and base their comment on lamprey on Coric et al. 2005. Our evidence for the presence of ASIC 3 in lamprey is that both the mechanical and pH response are blocked by APETx2 (Jalalvand et al., 2016). ASIC3 is present in both the peripheral and central nervous system in mammals.

In order to solve this issue, the authors could show expression of ASIC channels, and test the effect of a drug against ASIC1 on the response of somatostatin-expressing cells.

d) Finally, the only ASIC channel expressed in lamprey, ASIC1, when cloned and tested, has been reported to be proton insensitive (see https://www.ncbi.nlm.nih.gov/pmc/articles/PMC1464184/) – how then to explain then the activation of a subset of ciliated neurons exposed to protons?

It is correct (Coric et al. 2005) that the ASIC1 clone extracted from lamprey was not pH sensitive when expressed in frog oocytes, but our data show that the mechanical/pH response in the lamprey somatostatin expressing CSF cells in lamprey is mediated by ASIC3, and there is no evidence to suggest that ASIC3 should not be present in lamprey. The dopamine-containing cells are not pH-sensitive.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

[…] Specific comments from one reviewer:

The authors satisfactorily answered some of the issues raised on the quantification of images and provided high resolution images for their outstanding data acquired at high resolution. These data are very convincing and beautiful. Congratulations, this will be very precious and inspirational for further studies in the spinal field !

The issue remaining are related to the interpretation of the physiological results. The authors should either show expression of ASIC3 in CSF-cNs by ISH or tone down their conclusions relative to the ASIC3 receptor in response to pH.

We have reworded our text on page 8 and 13.

Similarly the authors should be more careful for stating the role PKD2L1 in pH sensing or mechanoreception as they lack genetic methods to investigate its function in lamprey.

Since the mechanosensitivity is mediated by an APETx2 sensitive mechanism in somatostatin CSF-c neurons but through another mechanism in the dopamine CSF-c neurons it is natural to point out the possibility of PKDL1, which is responsive in the zebrafish.

The text on page 8 now reads:

“As PKD2L1 has been confirmed as a mechanosensitive ion channel in zebrafish (Bohm et al., 2016, Sternberg et al., 2018, Orts-Del'Immagine et al., 2020), the results suggest that the mechano-sensitivity of the dopaminergic CSF-c neurons may be mediated by PKD2L1 channels.”

A) On the role of ASIC3 in pH sensing :

As stated in my first review, ASIC3 can only be found in the genome of mammals. Only ASIC1 has been identified in the lamprey genome (https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3047259) and the lamprey ASIC1a is proton insensitive (https://www.ncbi.nlm.nih.gov/pmc/articles/PMC1464184/).

The authors did not respond to my request of testing the role of ASIC1a in their experiments using pharmacology.

We have not tested ASIC1a blockers, since this channel in lamprey is known to not respond to pH (Coric et al. 2005), and ASIC1a is not blocked by APETx2.

One can therefore wonder whether 2uM APETx2 used by the authors (about 100X the EC50 of ASIC3 homomers, https://pubmed-ncbi-nlm-nih-gov/17113616/) could be acting on other targets (see suggested actions on Na or K channels mentioned here, https://pubmed-ncbi-nlm-nih-gov/17113616/). Please discuss clearly in the text.

We work within the specific dose range as described by Diochot et al. 2004, and the same range as has been used in mammalian dorsal root ganglion cells. We also note that APETx2 blocks (the ASIC3 blocker) the mechanosensitive response in somatostatin cell, whereas it has no effect on the mechanosensitivity of dopamine CSF-c cells. This strongly suggests that the effect of APETx2 is not a general unspecific effect on lamprey neurons.

We have added (the following text in the Discussion on ine page 13):

“The mechano-sensitivity of the dopamine and somatostatin expressing CSF-c neurons are mediated by different cellular mechanisms, the latter is blocked by APETx2, a selective ASIC3 blocker as shown in previous studies (E. Jalalvand, B. Robertson, H. Tostivint, et al., 2016; E. Jalalvand, B. Robertson, P. Wallen, et al., 2016). et al. et al. […] The data supports the previous interpretation that ASIC3 mediates both the acid-sensing and the mechanosensitivity in these CSF-c neurons.”

We also responded previously to this issue in our response:

In the article of Coric et al. 2005 they had identified one clone of cDNA that corresponded to ASIC1 and when expressed in oocytes, no pH sensitivity was found under these conditions. They do not comment regarding the possible presence of ASIC3. The review of Grunder and Chen (2010) has a focus on ASIC1a and base their comment in lamprey on Coric et al. 2005. Our evidence for the presence of ASIC3 in lamprey is that both the mechanical and pH response are blocked by APETx2, a selective antagonist of ASIC3, (Jalalvand et al. 2016, Nature. Com), strongly suggesting the presence of ASIC3 in the lamprey. ASIC3 is present in both the peripheral and central nervous system in mammals.

B) On the role of PKD2L1 in mechanoreception:

The authors summarize that PKD2L1 is responsible for mechanoreception only in dopaminergic CSF-cNs.

However:

1. PKD2L1 is expressed in all CSF-cNs, ventral (dopaminergic) and dorsal (somatostatinergic) as shown by the authors here and found in zebrafish (Djenoune 2014) and mouse (Huang 2006; Petracca 2018). In fact, in their 2016b publication, the authors had proposed that SST+ CSF-cNs were responding to basic pH via PKD2L1, suggesting a role for this channel in these cells.

Why therefore in this study, only mentioning the role of PKD2L1 for mechanoreception in the dopaminergic CSF-cNs despite the expression being there in both dorsal and ventral cell types?

Our results clearly show that there are different cellular mechanisms for the mechanotransduction, one blocked by APETx2 in somatostatin CSF-c cells and not in dopamine CSF-c neurons. Since PKD2L1 is present in the dopamine cells and known to be mediate mechanosensitivity in zebrafish CSF-c cells it is natural to point to this possibility.

2. Due to the lack of specific antagonists, the authors do not have the tools in lamprey to measure its contribution to mechanoreception. They can only suggest that PKD2L1 contribute to mechanoreception in both ventral and dorsal CSF-cN types.

The mechanosensitive mechanisms in the two types of CSF-c cells, the laterally projecting and ventral CSF-c neurons, in lamprey must be different, given the difference in APETx2 effects.

3. Please correct an error in citation and references used:

The authors use Bohm et al. 2016 to state that CSF-cNs rely on PKD2L1 to be mechanosensory in zebrafish.

To be correct, we showed in vivo in Bohm 2016 that both ventral and dorsal CSF-cNs respond to concave (not convex) mechanical deformations of the spinal cord via PKD2L1 (response are abolished in the KO).

However, mechanoreception cannot be rigorously demonstrated in vivo. It is therefore only in Sternberg 2018 that we could show in vitro using a piezo device to mechanically stimulate their membrane that all CSF-cNs isolated in primary cultures are mechanosensory cells and that their response always rely on PKD2L1.

Note that in zebrafish, we now understand that CSF-cNs in vitro do not respond to CSF flow (Prendergast et al. under review) and in vivo, their response to concave mechanical bending of the spinal cord needs their interaction with the Reissner fiber (Orts Dell Immagine 2020), which does not alter itself the flow (Cantaut-Belarif 2018).

Interesting with the results with Reissner´s fiber – a possibility that has been considered often but without substantial evidence previously, but not citable as yet.

The authors should at minima cite Sternberg et al. 2018 for showing the role of PKD2L1 in mechanoreception, but to be fair, also propose with more nuances in the discussion how these cells in lamprey can combine ASIC and PKD channels to sense pH and mechanical inputs, citing Bohm 2016, Sternberg 2018 and Orts Del Immagine 2020.

Done.

https://doi.org/10.7554/eLife.73114.sa2

Article and author information

Author details

Elham Jalalvand

Department of Applied Physics and Science for Life Laboratory, KTH Royal Institute of Technology, Stockholm, Sweden

Contribution
Data curation, Formal analysis, Investigation, Methodology, Validation, Visualization, Writing - original draft, Writing - review and editing

Competing interests
No competing interests declared

"This ORCID iD identifies the author of this article:" 0000-0003-4759-3301
Jonatan Alvelid

Department of Applied Physics and Science for Life Laboratory, KTH Royal Institute of Technology, Stockholm, Sweden

Contribution
Formal analysis, Methodology, Writing - review and editing

Competing interests
No competing interests declared

"This ORCID iD identifies the author of this article:" 0000-0002-3554-9322
Giovanna Coceano

Department of Applied Physics and Science for Life Laboratory, KTH Royal Institute of Technology, Stockholm, Sweden

Contribution
Formal analysis, Methodology, Writing - review and editing

Competing interests
No competing interests declared
Steven Edwards

Department of Applied Physics and Science for Life Laboratory, KTH Royal Institute of Technology, Stockholm, Sweden

Contribution
Formal analysis, Methodology, Writing - review and editing

Competing interests
No competing interests declared
Brita Robertson

Department of Neuroscience, Karolinska Institutet, Stockholm, Sweden

Contribution
Formal analysis, Writing - review and editing

Competing interests
No competing interests declared
Sten Grillner

Department of Neuroscience, Karolinska Institutet, Stockholm, Sweden

Contribution
Writing - review and editing

Competing interests
No competing interests declared

"This ORCID iD identifies the author of this article:" 0000-0002-8951-3691
Ilaria Testa

Department of Applied Physics and Science for Life Laboratory, KTH Royal Institute of Technology, Stockholm, Sweden

Contribution
Methodology, Supervision, Validation, Visualization, Writing - original draft, Writing - review and editing

For correspondence
ilaria.testa@scilifelab.se

Competing interests
Reviewing Editor, eLife

"This ORCID iD identifies the author of this article:" 0000-0003-4005-4997

Funding

Swedish Foundation for Strategic Research (FFL15-0031)

Ilaria Testa

Swedish Research Council (2021-01995)

Sten Grillner

Royal Institute of Technology (Göran Gustafssons Stiftelse)

Ilaria Testa
Elham Jalalvand

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

Acknowledgements

We thank the ALM facility at the Science for Life Laboratory for access to the STED Leica Sp8. We thank the SciLifeLab SFO funding, the GGS foundation, and the Swedish Foundation for Strategic Research for supporting the project FFL15-0031, and grants from The Swedish Research Council to SG (2021-01995).

Ethics

All experiments were performed in accordance with animal welfare guidelines set forth by Karolinska Institutet and were approved by Stockholm North Ethical Evaluation Board for Animal Research.

Senior and Reviewing Editor

Suzanne R Pfeffer, Stanford University School of Medicine, United States

Reviewers

Francesca Bottanelli
Claire Wyart, Institut du Cerveau et la Moelle épinière, Hôpital Pitié-Salpêtrière, Sorbonne Universités, UPMC Univ Paris 06, Inserm, CNRS, France

Publication history

Preprint posted: August 17, 2021 (view preprint)
Received: August 17, 2021
Accepted: January 19, 2022
Version of Record published: February 1, 2022 (version 1)

Copyright

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.