Research Articles: Systems/Circuits

Tilt In Place Microscopy (TIPM): a simple, lowcost solution to image neural responses tobody rotations
https://doi.org/10.1523/JNEUROSCI.1736-22.2022
Cite as: J. Neurosci 2022; 10.1523/JNEUROSCI.1736-22.2022
Received: 13 September 2022
Revised: 23 November 2022
Accepted: 7 December 2022

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Tilt In Place Microscopy (TIPM): a simple,
low-cost solution to image neural responses tobody rotations
Kyla R. Hamling1, Yunlu Zhu1, Franziska Auer1, and David Schoppik1
1 Depts. of Otolaryngology, Neuroscience & Physiology, and the Neuroscience Institute, New York University Grossman

School of Medicine, New York, NY 10016

1

Abstract2

Animals use information about gravity and other destabilizing forces to balance and3navigate through their environment. Measuring how brains respond to these forces4

requires considerable technical knowledge and/or ﬁnancial resources. We present a5

simple alternative: Tilt In Place Microscopy (TIPM). TIPM is a low-cost and non-6

invasive way to measure neural activity following rapid changes in body orientation.

7

Here we used TIPM to study vestibulospinal neurons in larval zebraﬁsh during and8

immediately after roll tilts. Vestibulospinal neurons responded with reliable increases

9in activity that varied as a function of ipsilateral tilt amplitude. TIPM diﬀerentiated10

tonic (i.e. sustained tilt) from phasic responses, revealing coarse topography of stim-

11

ulus sensitivity in the lateral vestibular nucleus. Neuronal variability across repeated12

sessions was minor relative to trial-to-trial variability, allowing us to use TIPM for

13

longitudinal studies of the same neurons across two developmental timepoints. There,

14

we observed global increases in response strength, and systematic changes in the neu-

15

ral representation of stimulus direction. Our data extend classical characterization of16

the body tilt representation by vestibulospinal neurons and establish TIPM’s utility to17

study the neural basis of balance, especially in developing animals.

18

Signiﬁcance Statement

19 Vestibular sensation inﬂuences everything from navigation to interoception. Here we detail
20 a straightforward, validated and nearly-universal approach to image how the nervous system

21 senses and responds to body tilts. We use our new method to replicate and expand upon
22 past ﬁndings of tilt sensing by a conserved population of spinal-projecting vestibular neurons.
23 The simplicity and broad compatibility of our approach will democratize the study of the
24 brain’s response to destabilization, particularly across development.

25

Introduction

26 Animals must transform forces acting on the head/body into commands to stabilize gaze/posture,
27 orient, navigate, and regulate physiology (Angelaki and Laurens, 2020; Chen et al., 2021;
28 Daltorio and Fox, 2018; Yates et al., 2013; Yoder and Taube, 2014). Both sensory organs and29 bodies change as animals develop and age. Studying neuronal representations of forces – par30 ticularly over time – presents a profound challenge: to measure brain activity while applying
31 destabilizing forces. For over a century novel apparatus have met this challenge, primarily
32 in studies of the vertebrate vestibular system: from observational chambers (Lowenstein and
33 Roberts, 1949; Mach, 1886; Straka et al., 2021), translating sleds (Fleisch, 1922), and later
34 to electrophysiology-compatible rotating swings (Adrian, 1943), modern platforms with 6
35 degrees of freedom (Angelaki et al., 1999; Branoner and Straka, 2018) and wireless recording
36 (Carriot et al., 2017). Forces are usually delivered sinusoidally, conferring mechanical advan37 tages and facilitating linear systems analysis and modelling of neuronal/behavioral responses
38 (Laurens et al., 2017). While studies of vestibular processing have advanced nearly every
39 area of systems neuroscience (Goldberg et al., 2012a), each apparatus and stimulus paradigm
40 represents a set of necessary trade-oﬀs. The specialized hardware used in most experiments
41 is both a cost and knowledge barrier. Further, few existing approaches permit repeated
42 measures of activity from the same neurons over days, key to understanding changes that43 accompany development (Beraneck et al., 2014; Peusner, 2014) and learning (Goldberg et
44 al., 2012b). Here we detail and validate an approach for longitudinal measures of vestibular
45 sensitivity.

46

Imaging neuronal activity by measuring changes in ﬂuorescence of genetically-encoded

47 calcium indicators has transformed neuroscience. The vestibular ﬁeld has been comparatively
48 slow to adopt this technology largely because imaging neurons while animals are challenged
49 with vestibular stimuli is technically demanding. Recent eﬀorts include microscopes that50 rotate (Migault et al., 2018), image a rotating sample (Hennestad et al., 2021; Tanimoto et al.,
51 2022), and optically trap and move part of the vestibular apparatus (Favre-Bulle et al., 2018).
52 Taken together, these studies clearly illustrate the promise of using modern microscopy and
53 genetics to understand the vestibular system. However, each requires specialized optical
54 and/or engineering expertise to implement, limiting their impact. Motivated by these studies,
55 we sought to develop a complementary low-cost, straightforward means to image neural
56 activity following body tilts.
57

We focused our eﬀorts on a model vertebrate with a rich vestibular repertoire: the larval

58 zebraﬁsh. At four days old larval zebraﬁsh swim freely and maintain posture and stabilize
59 gaze (Bagnall and Schoppik, 2018). Both behaviors require central vestibular neuronal cir60 cuits (Schoppik et al., 2017), with considerable development between four and seven days
61 (Bianco et al., 2012; Ehrlich and Schoppik, 2017, 2019). Importantly, the larval zebraﬁsh
62 is both transparent and genetically-accessible, facilitating measurements of calcium indica63 tors in vestibular neurons, and/or loss-of-function assays in mutants (Whitﬁeld et al., 1996).
64 Finally, like all vertebrates, the larval zebraﬁsh has a small population of vestibulospinal neu65 rons (˜50 cells) with reliable and well-characterized responses to tilt stimulation (Hamling
66 et al., 2021; Liu et al., 2020). Taken together, the zebraﬁsh is a powerful model to develop
67 new methods to investigate vestibular function.
68

Here we present data from a new approach to imaging neural activity in response to body69 rotations we call Tilt In Place Microscopy, or TIPM. TIPM allows extremely rapid (<6 ms)
70 whole-body rotations toward and away from the horizon, allowing precise characterization of
71 tilt sensitivity. We validated TIPM by characterizing the roll responses, topography, and de72 velopment of larval zebraﬁsh vestibulospinal neurons. We found that vestibulospinal neurons

73 respond reliably to ipsilateral steps with parametrically increasing activity, consistent with74 prior electrophysiological measurements in ﬁsh and mammals (Peterson, 1970; Rovainen,
75 1979). We repeated TIPM sequentially on the same ﬁsh and found that trial-to-trial vari76 ation was likely intrinsic to vestibulospinal responses, not due to our approach/apparatus.
77 Vestibulospinal neurons had a comparatively small response to phasic stimulation; neurons
78 that sensed phasic components were preferentially located in the ventral lateral vestibular
79 nucleus. The bulk of the vestibulospinal response was derived from utricular sensation. Fi80 nally we measured responses from the same neurons at two behaviorally-relevant time points,
81 revealing increased response strength in older neurons and systematic changes in directional
82 selectivity as neurons develop. We end with a discussion of the advantages of our method
83 (low cost, broad compatibility, extensibility) and its limitations. Our method will facilitate
84 investigation of neuronal responses to tilt stimulation, particularly in small model animals
85 such as Drosophila, Caenorhabditis, Danio, Danionella, and Xenopus.

86

Materials and Methods

87

Fish Care

88 All procedures involving zebraﬁsh larvae (Danio rerio) were approved by the Institutional
89 Animal Care and Use Committee of New York University Grossman School of Medicine.
90 Fertilized eggs were collected and maintained at 28.5°C on a standard 14/10 hour light/dark
91 cycle. All experiments were performed on larvae between 4 and 7 dpf. During this time,
92 zebraﬁsh larvae have not yet diﬀerentiated their sex into male/female. Before 5 dpf, larvae
93 were maintained at densities of 20-50 larvae per petri dish of 10 cm diameter, ﬁlled with
94 25-40 mL E3 with 0.5 ppm methylene blue. After 5 dpf, larvae were maintained at densities
95 under 20 larvae per petri dish in E3 and were fed cultured rotifers (Reed Mariculture) daily.

96

Fish Lines

97 All experiments were done on the mitfa -/- background to remove pigment for
98 imaging.

All larvae were labeled with Tg(nefma:GAL4;UAS:GCaMP6s) (Liu et al.,

99 2020; Thiele et al., 2014).

For experiments testing utricular origin of responses,

100 Tg(nefma:GAL4;UAS:GCaMP6s) ﬁsh with a homozygous recessive loss-of-function muta101 tion of the inner ear-restricted gene, otogelin (otog-/-), also called rock solo AN66 (Whitﬁeld
102 et al., 1996) were visually identiﬁed by a bilateral lack of utricular otoliths.

103

Imaging Set-Up and Apparatus

104 Larval ﬁsh (4-7 dpf) were mounted in a small drop of 2% low melting point agarose (Ther105 moFisher Scientiﬁc 16520) in the center of the uncoated side of a mirror galvanometer (Thor106 labs GVS0111). E3 was placed over the agarose and the galvanometer mirror was placed
107 under the microscope.
108

For routine imaging, a baseline voltage was applied to the galvanometer to set it at one

109 end of its range, allowing for up to 40° of rotation away from the horizontal plane in one
110 direction. Stimuli were capped at ±30° to allow the experimenter to apply an additional
111 small oﬀset voltage to correct for slight deviations from horizontal incurred while mounting
112 the ﬁsh, and because of steric limitations relative to the objective. All trials (3 trials per
113 step size, 2 stimuli repeats per trial) in one direction were run with that baseline voltage
114 manually set at horizontal, then a new baseline voltage was applied and the galvanometer
115 was re-centered at horizontal to continue performing trials in the opposite tilt direction.
116 Experiments with rock solo ﬁsh were performed with a diﬀerent stimulus paradigm; in these
117 experiments no baseline voltage was applied to the galvanometer before it was positioned at
118 horizontal, and the maximum voltage drive was then applied during the stimulus to rotate
119 the sample to ±20°.
120

For experiments where responses at horizontal were compared to responses measured

121 directly at the eccentric angle, ﬁsh were mounted and ﬁrst imaged upon return to horizontal.
122 Then, the microscope was manually focused to allow the ﬁsh to be in focus at an eccentric
123 angle for subsequent trials. Fish were then anesthetized by applying 0.2 mg/mL ethyl124 3-aminobenzoic acid ethyl ester (MESAB, Sigma-Aldrich E10521) to the ﬁsh mounted in
125 agarose. The ﬁsh was allowed to sit with the anesthetic for 10 minutes before imaging
126 recommenced to ensure it had reached full eﬀect; MESAB remained on the ﬁsh for the rest
127 of the imaging session to keep the ﬁsh properly anesthetized. The imaging was then repeated
128 in the anesthetized ﬁsh with the ﬁsh in focus upon return to horizontal and subsequently in
129 focus at the eccentric angle.
130

Galvanometer control was done in Matlab 2019b (Mathworks, MA) using the Data Acqui-

131 sition Toolbox to interface with a data acquisition card (PCIe-6363, National Instruments,
132 TX). Custom code was written to simultaneously (1) deliver an analog waveform to control
133 the galvanometer (2) measure the analog voltage that corresponded to the galvanometer po134 sition and (3) deliver synchronizing digital pulses to begin imaging. For all step and impulse
135 stimuli, the galvanometer was allowed to step to/away the eccentric angle at the maximum
136 angular velocity/acceleration achievable (Table 1). Impulse stimuli were delivered in both
137 directions. A microscope (Thorlabs Bergamo) was used to measure ﬂuorescence elicited by
138 multiphoton excitation (920 nm) from a pulsed infrared laser (MaiTai HP, Newport, CA).
139 Fast volumetric scanning was achieved using a piezo actuator (ThorLabs PFM450E) to move
140 the objective. Each frame of the volume (416 x 64 pixels) was collected with a 1 μs pixel
141 dwell time (18.6 frames/second). Volumes ranged from 6-9 planes in depth (6 μm between
142 planes) to cover the entire vestibulospinal nucleus, resulting in a mean eﬀective volume rate
143 of 2.2 volumes per second (range 1.9-2.7 volumes/second). Fish that were imaged multiple
144 times were imaged using the same scan settings across both sessions. The rock solo mutant
145 ﬁsh and their wild-type siblings were imaged prior to the addition of the piezo actuator; in
146 the place of volumetric imaging, for these experiments single z-planes were imaged separately
147 (2-3 planes per ﬁsh) at a frame rate of 6.6 frames/second.

148

Data Analysis & Statistics

149 Calcium response extraction and analysis
150 Data analysis was performed using custom code in Matlab 2017b (Mathworks, MA). We
151 pre-processed the imaging data with code adapted from the CalmAn package (Giovannucci
152 et al., 2019), and then performed NoRMCorre rigid motion correction (Pnevmatikakis and
153 Giovannucci, 2017) on our GCaMP6s signal across all concatenated trials of the same stim154 ulus type for each ﬁsh. We then hand-drew polygon ROIs in FIJI (Schindelin et al., 2012)155 around each vestibulospinal cell on the maximum intensity projection of all the motion cor156 rected frames. We imported ROIs into Matlab using ReadImageJROI (Muir and Kampa,
157 2015) and extracted the mean pixel value across each ROI for all timepoints of each trial,
158 and performed normalization of the raw ﬂuorescence trace.
159

For experiments imaged at horizontal, we quantiﬁed the peak calcium response to each

160 stimulus as the mean ΔF/F in the ﬁrst second after the sample returned to horizontal. For
161 experiments imaged at the eccentric angle, responses calculated for comparison to those at
162 horizontal were the mean ΔF/F in the last second before the sample returned to horizontal.
163 A cell was determined to have a signiﬁcant response to a stimulus if the peak calcium
164 responses across all trials were signiﬁcantly higher (one-tailed t-test, p¡0.05) than the mean
165 calcium responses during the ﬁrst second of the baseline period of that same cell. For each
166 cell, a directionality index was calculated by taking the diﬀerence between the peak calcium
167 response to ipsilateral and contralateral 30° steps, normalized by their sum.
168

To calculate the sensitivity of peak calcium responses to step magnitude, we ﬁt a line

169 with two free parameters to the peak calcium responses from all trials of all step magnitudes
170 in a single direction (ipsilateral or contralateral). During analyses of longitudinal calcium
171 imaging, cells with a signiﬁcant sensitivity increase/decrease between time points was deﬁned
172 as follows: To determine a cutoﬀ for signiﬁcantly increasing/decreasing slopes, for each ﬁsh
173 we shuﬄed the peak calcium responses across all trials from both ages in the same direction,

174 and used the shuﬄed responses to calculate a best-ﬁt line with a slope. We then took the
175 diﬀerence of the shuﬄed slopes between 4 and 7 dpf for each cell for the ipsilateral and
176 contralateral direction. The cutoﬀ for signiﬁcant sensitivity change was deﬁned as the mean
177 of the shuﬄed slopes ± 2 SD. In longitudinal experiments, “Early Contra Responders” were
178 deﬁned as cells that had a contralateral slope at 4 dpf greater than the mean + 2 SD of
179 contralateral 4 dpf shuﬄed slopes.
180

To determine any ﬁeld-of-view shifts after an eccentric step, we performed a two-

181 dimensional normalized cross-correlation between the frame prior to the stimulus and the
182 frame after the stimulus for each plane of the volume. These analyses were performed on
183 frames from unprocessed volumes. Field-of-view shift in the x- and y-axis was determined by
184 ﬁnding the position of highest correlation coeﬃcient within the resulting matrix, and corre185 sponding that matrix with an x- and y-axis shift in pixels (reported in text after conversion
186 to μm). Mean correlation coeﬃcients for each ﬁsh were calculated from the center of the
187 correlation coeﬃcient matrix (i.e. the correlation between the two frames without any x- or
188 y-shift).
189

Correlation was evaluated using Pearson’s correlation coeﬃcient (ρ). We report slope ﬁts

190 and 95% conﬁdence intervals (CI). To test for interactions between groups we use either a
191 univariate analysis of variance test (ANOVA) or multivariate (MANOVA).

192 Normalization
193 When comparing activity in the same neuron measured at horizontal only (our standard
194 imaging paradigm), we normalized the ﬂuorescence against the mean ﬂuorescence value in
195 the last 5 seconds of the baseline period within each trial. When comparing activity in the
196 same neuron measured at diﬀerent angles, we used the ﬂuorescence measured in an anes197 thetized condition at the angle at which the imaging was done. Our rationale is as follows:
198 the intensity of a genetically-encoded calcium indicator reﬂects a number of variables, neces199 sitating normalization. The “baseline” level is usually derived during a neutral condition with

200 respect to the stimulus, correcting for diﬀerences in expression levels and variable imaging
201 conditions (e.g. IR light penetration or scattering of emitted photons). Further, vestibular
202 neurons might have diﬀerent basal activity when held at eccentric positions. We assert that
203 the ﬂuorescent intensity measured in a given neuron in an anesthetized animal will only
204 reﬂect basal expression levels and variability due to imaging conditions. Consequentially,
205 it is an easily-accessible baseline that permits us to compare responses in the same neuron
206 when held at diﬀerent angles.

207 Vestibulospinal Cell Body Position and Roll Angle Analysis
208 For each ﬁsh, an additional 2-photon volumetric stack was taken with scan settings optimized
209 for a high-signal, low-speed image (2 μs scan speed, cumulative averaging across 4 frames,
210 1 μm between z-planes). This stack was used for localizing the vestibulospinal cell bodies
211 in three-dimensions, deﬁned relative to the Mauthner cell lateral dendrite. To deﬁne these212 XYZ positions, we ﬁrst placed reference point ROIs in FIJI at the lateral-most tip of the
213 Mauthner cell lateral dendrite in both brain hemispheres. We then dropped point ROIs on
214 all vestibulospinal cells that were analyzed in our calcium imaging trials, placing the ROI
215 at the center of the cell body at the z-plane where the cell was most in-focus. Using FIJI’s
216 “Measure” tool, we measured the XYZ position of each ROI in microns, and exported this217 data to Excel. For each vestibulospinal cell position, we subtracted oﬀ the XYZ position
218 of the Mauthner lateral dendrite in its corresponding brain hemisphere to convert absolute
219 position to relative position, and then imported the relative XYZ position data to Matlab
220 for plotting.
221

For calculating the roll tilt angle of each ﬁsh, we used the left and right Mauthner lateral

222 dendrite reference ROIs to ﬁnd the distance between the two hemispheres in depth (z-axis).
223 We then calculated the average mediolateral (x-axis) distance between the Mauthner lateral
224 dendrites (171.8 μm). We took the arctangent of the z-distance and average x-distance to
225 calculate the roll angle for each mounting.

226

Results

227

Rationale, apparatus, and stimulus for Tilt In Place Microscopy

228 We developed a simple method (TIPM) to permit imaging of neuronal activity following
229 body tilts. The vestibular end-organs in vertebrates detect either linear accelerations (such
230 as orientation relative to gravity) or rotational accelerations. We reasoned that the most
231 straightforward way to assay this activity would be to image the same volume before and232 after such stimulation to avoid image registration challenges and to maximize compatibility
233 with diﬀerent microscope architectures. The kinetics of ﬂuorescent indicators of neuronal
234 activity are slow to decay (Chen et al., 2013). Consequentially, a suﬃciently rapid step back235 to the horizon from an eccentric orientation would produce a response with two components.
236 The ﬁrst component would reﬂect the steady-state activity of the neurons encoding linear
237 acceleration due to gravity (i.e. the body’s tilt) before the step. The second component
238 would reﬂect the phasic response to the step itself, if any. We refer to the stimulus that239 elicits these combined responses as a “step.” Complementarily, a second stimulus comprised
240 of a rapid step to an eccentric angle and back would generate a response to an impulse of
241 rotation, devoid of any steady-state component. We refer to this stimulus as an “impulse.”
242 Taken together, step and impulse stimuli allow for characterization of both the tonic (i.e.
243 steady-state body tilt) and phasic (i.e. rapidly changing) components of a neuron’s response.
244 The key to our method is therefore delivery of rapid rotations to the preparation.
245

Here we used a mirror galvanometer as a platform to rapidly rotate an immobilized larval

246 zebraﬁsh. Mirror galvanometers are the tool of choice to steer light to particular angles for247 their precision and rapid response. We mounted a larval zebraﬁsh in a small drop of low
248 melting temperature agarose directly on the uncoated side of the mirror (Figure 1A). We
249 could rotate the platform through nearly the full range of the galvanometer both rapidly
250 (5.3 ms for a 30° step, (Table 1), and precisely (Figure 1B-1D).
251

To image ﬂuorescence, the platform is mounted underneath a water-dipping objective

252 on a two-photon microscope capable of rapid volumetric scanning. All experiments were
253 conducted in complete darkness. A drop of water, held in place by surface tension, sits
254 between the agarose and the objective. Prior to starting an experiment, the platform is
255 adjusted to sit horizontally underneath the objective such that the neurons of interest are
256 in focus. We measured ﬂuorescence across the volume to deﬁne a baseline for the neurons.
257 For the step stimuli, we rotated the platform to an eccentric angle (where the neurons of
258 interest are no longer in focus), held the platform at that orientation for 15 seconds, and259 then returned the platform back to horizontal (Figure 1C). There, we measured the changes
260 in ﬂuorescent intensity in response to our stimulus.

261

Vestibulospinal neurons respond reliably to ipsilateral step stimuli

262 We began by measuring responses to roll tilts at 4 days post-fertilization (dpf) in vestibu263 lospinal neurons (Figure 1E) labelled in transgenic line, Tg(nefma:GAL4;14xUAS:GCaMP6s)
264 (Liu et al., 2020). Previous work in larval zebraﬁsh (Liu et al., 2020) and other animals (Fu265 jita et al., 1968) established that vestibulospinal neurons increase their activity as a function
266 of roll tilt angle, with a strong preference for tilts in the direction of the recorded neuron
267 (i.e. a cell in the left hemisphere responds when the left ear is rolled down; henceforth
268 called “ipsilateral roll”). Vestibulospinal neurons therefore provide an excellent test-bed to
269 evaluate new tilt paradigms, such as the step and impulse stimuli we use here.
270

We mounted the ﬁsh parallel to the platform’s axis of rotation to provide both ipsilat-

271 eral and contralateral roll tilts of 10°, 20°, and 30° (Figure 1C). We detected signiﬁcant
272 changes in ﬂuorescence relative to each cell’s own baseline (henceforth called “responsive
273 cells”, Methods) in 94% of neurons (67/71 neurons from 10 ﬁsh) (Figure 1F, example re274 sponsive trace). Importantly, responses were reliable across repeated trials, with a median
275 coeﬃcient of variation across trials of 0.19 (Figure 1F,1G). Consistent with prior reports,
276 responses were direction-dependent. The majority of neurons had a larger response to ipsi277 lateral roll compared to contralateral (Directionality Index = 0.46 ± 0.43) (Figure 1H,1J).

278 Additionally, we found that the strength of responses to roll stimuli increased with the size
279 of the step. For steps in both the ipsilateral and contralateral direction, we observed that
280 the peak response, deﬁned as the mean response within the ﬁrst second after the end of
281 the stimulus, scaled linearly with the magnitude of the roll step (Figure 1H,1I) (mean slope
282 of responsive neurons= 0.07 ± 0.06 ΔF/F/° ipsilateral, 0.02 ± 0.02 ΔF/F/° contralateral).
283 We conclude that our apparatus can elicit reliable, parametric, and directionally-sensitive
284 responses following roll tilts of diﬀerent amplitude in vestibulospinal neurons.

285

TIPM is robust to extrinsic sources of variability

286 There are several potential sources of variability that could compromise detection of reliable
287 ﬂuorescent changes following stimulation. First we measured response variation from the fol288 lowing sources: (1) ﬁeld-of-view movement during imaging and (2) mounting variability. We
289 then measured intrinsic trial-to-trial variability that presumably reﬂects biological sources
290 such as changes in intraneuronal calcium, spike rate ﬂuctuations, or state of the animal
291 (Schoppik et al., 2008). If the variability observed from trial-to-trial is greater than extrinsic
292 variability, we would conclude that our approach is suﬃciently robust.
293

The dynamic nature of TIPM introduces the potential for the ﬁeld-of-view of our sample

294 to move during the course of imaging. Sample movement has the potential to cause variability
295 in ﬂuorescence intensity that does not reﬂect an underlying calcium ﬂuctuation. Qualita296 tively, we did not observe ﬁeld-of-view shifts acutely between the baseline recording period
297 and after an eccentric step. We quantiﬁed such changes by performing a cross-correlation of
298 each frame before and after the eccentric step. To eliminate signal changes from neuronal299 ﬂuctuations, we performed this analysis on unprocessed volumes measured in anesthetized
300 ﬁsh. The frames before and after the eccentric step were most-highly correlated with each
301 other when they were not shifted relative to each other (mean shift = 0.02 μm in x-axis,
302 -0.01 μm in y-axis; mean correlation without shift = 0.5). Additionally the mean peak ﬂuo303 rescence change after the eccentric step in anesthetized ﬁsh was very low (0.08±0.09 ΔF/F304 anesthetized vs 2.2±1.8 ΔF/F un-anesthetized, n=26 neurons), indicating there is very lit305 tle variation in the ﬂuorescence signal that results from acute shifts during imaging. We
306 conclude that TIPM as implemented here introduces tolerable levels of sample movement.
307

Notably, as with any long-term imaging experiment, we did observe that some samples

308 slowly drift in X/Y/Z between trials. We estimate this drift at approximately 1 μm/minute309 in all axes. These slow shifts can be easily corrected either manually between trials or by
310 post-hoc motion correction and so do not introduce appreciable variability into measured
311 responses.
312

We next addressed the variability due to mounting. For each imaging experiment, larval

313 ﬁsh are manually mounted on the galvanometer in agarose in a dorsal-up position. Every
314 attempt is made to minimize roll, pitch, and yaw relative to the axis of rotation, but manual
315 mounting is subject to small variations. These variations would impose linear accelerations
316 that scale with the distance from the center of the axis of rotation. Such shifts would be
317 challenging to quantify and, if large, might compromise longitudinal experiments.
318

To estimate how much variation in response originated from variation in mounting, we

319 performed a repeated imaging experiment. We mounted and imaged a ﬁsh, then removed the
320 ﬁsh from agarose and re-mounted the same ﬁsh and repeated our imaging protocol (Figure
321 2A). We were able to reliably identify neurons between the ﬁrst and second mounts (Figure322 2B). To estimate the roll tilt, we calculated the bilateral diﬀerence in z-position of the tips
323 of the left and right Mauthner lateral dendrites, and used this to calculate a roll angle of the
324 head. We observed only minor rotation of the baseline position of the ﬁsh in the roll axis
325 between the ﬁrst and second mounts (2.4±1.0°, N=5 ﬁsh).
326

We saw a strong correlation (ρ=0.83) in the response of individual neurons between the

327 ﬁrst and second mount (Figure 2C-2D). We did not observe a signiﬁcant shift in responses
328 between the ﬁrst and second mounts (paired t-test p=0.64, n=34 neurons, N=5 ﬁsh), and
329 the responses of the neurons fell nearly along the unity line (slope=0.75 ± 0.18 CI).
330

To contextualize the magnitude of the mount-to-mount variability we observed, we com-

331 pared it to trial-to-trial variability. Response correlation between TIPM sessions is compa332 rable to the response correlation between two subsequent trials (Figure 2E) within a single
333 imaging session (ρ=0.79, slope=0.79 ± 0.09 CI, n=34 cells, N=5 ﬁsh). These data suggest
334 that most of the variability in response magnitude we see mount-to-mount reﬂects inherent
335 variability.
336

Together these experiments establish “best practices” to estimate variability when using

337 TIPM to measure neural responses. We conclude that variability due to our apparatus or
338 mounting are relatively minor concerns for estimating neural response magnitude in our
339 preparation.

340

Neural activity imaged after a step reﬂects the encoding of body341

tilt prior to the step

342 Our interpretation of the response rests on the assumption that the activity observed after
343 the platform returns to the horizon primarily reﬂects the activity of the neuron at the
344 eccentric position. To test this assumption, we compared the response of neurons at eccentric
345 angles to that after a step returning the ﬁsh to horizontal. In this experiment, we presented
346 the same 30° roll stimulus to ﬁsh while measuring activity ﬁrst upon return to the horizontal
347 plane as previously described (Figure 3A, black), then on subsequent trials measuring activity
348 directly at the eccentric 30° angle (Figure 3A, magenta). Because the light path to the neuron
349 changes as a function of eccentricity, we normalized ﬂuorescence to a baseline stack taken
350 at either the horizontal or the eccentric angle while the ﬁsh was anesthetized. We compared
351 ﬂuorescence in vestibulospinal neurons in the ﬁrst second upon return to horizontal to the352 responses of the same neurons in the last second of the eccentric step. Neural responses at
353 eccentric angles were closely correlated (ρ=0.94) with the responses measured at horizontal
354 (Figure 3B,3C). The strong similarity in response amplitude supports two conclusions: First,
355 that the response of the neuron upon return to the horizontal is indeed a reasonable proxy356 for a neuron’s activity at an eccentric angle in the moment just prior to the return step.
357 Second, by inference, larval zebraﬁsh vestibulospinal neurons should have comparatively
358 small responses to phasic stimulation.
359

Notably, while measuring ﬂuorescent intensity at the eccentric angle, we observed quite

360 diﬀerent dynamics among vestibulospinal neurons. The most striking variation was in the
361 decay-rate of the ﬂuorescent intensity (Figure 3D). Some neurons had a distinct peak fol362 lowed by a fast-decay (Figure 3B, left), while others had plateau-like responses that had
363 little to no decay over the 15 second hold (Figure 3B, right). In all neurons measured, ﬂuo364 rescent intensity reached its peak within 10 seconds of being at the eccentric angle (median
365 5.2 s) (Figure 3E). We conclude that for vestibulospinal neurons a 10 second step would be
366 suﬃcient to ensure accurate detection of the peak response upon return to horizontal (cor367 relation between eccentric calcium response at its peak and at 10 seconds, ρ=0.91). As this368 value will vary between neuronal populations, preliminary experiments to set the optimal
369 window should be done for each new population of interest. By adjusting the length of the
370 TIPM eccentric step to one’s own experimental goals and observed calcium dynamics, the371 experimenter can use the return to horizontal response as a proxy to measure the magnitude
372 of either the peak or steady-state calcium responses. Taken together, measuring ﬂuores373 cence upon return to horizontal can be used to accurately extrapolate information about
374 the neuron’s response at the eccentric angle. Further, while imaging at horizontal alone can
375 not provide information about temporal dynamics, we demonstrate here how TIPM can be
376 modiﬁed to allow for imaging at an eccentric angle to study variations in response dynamics
377 within a population of neurons.

378

Vestibulospinal neurons respond weakly to impulses of angular ac-379

celeration

380 To measure the impulse response of vestibulospinal neurons, we delivered rapid roll steps to381 10°, 15°, or 30° and then back to horizontal in <13 ms (Figure 4A,4B; Table 1). We observed
382 signiﬁcant changes in ﬂuorescence to the impulse stimulus in a moderate fraction (35.4%)
383 of neurons (n=22/62 neurons from N=6 ﬁsh). The average peak ﬂuorescence observed to
384 the impulse stimulus was small (0.53 ± 0.39 ΔF/F for an ipsilateral 30° stimulus) compared
385 to the response to the tilt stimulus. Impulse responses are more variable across trials than
386 responses to the step stimuli (median coeﬃcient of variation = 0.76 vs 0.19) (Figure 4D).
387 Unlike responses to steps of diﬀerent amplitudes, peak ﬂuorescent intensity did not vary
388 systematically with the magnitude of the impulse (slope of peak ﬂuorescence = 0.002 ± 0.01
389 ΔF/F/° ipsilateral, -0.002 ± 0.02 ΔF/F/° contralateral) (Figure 4F). Additionally, impulse
390 responses did not show a consistent directional-preference. Most neurons responded equally
391 strongly to ipsilateral and contralateral steps (Directionality Index = 0.10 ± 0.37) (Figure392 4E,4G).
393

We asked whether there was topographical organization to these responsive neurons

394 within the lateral vestibular nucleus. While non-responsive neurons are found distributed
395 evenly throughout the lateral vestibular nucleus, neurons with a signiﬁcant response to the
396 impulse stimulus are located more ventro-laterally (mean dorsoventral position relative to397 Mauthner lateral dendrite = -3.6±7.0 μm responsive neurons vs 2.2±11.0 μm non-responsive
398 neurons; p=0.03) (Figure 4H,4I). There may therefore be topographic diﬀerences in inner399 vation by VIIIth nerve aﬀerents that relay phasic vestibular inputs relative to tonic inputs,
400 such that impulse-responsive aﬀerents target only a subset of ventro-lateral vestibulospinal
401 neurons.
402

Taken together, our data argue that vestibulospinal neurons are more sensitive to the403 tonic component of the step stimulus than to an impulse stimulus. We infer that the response404 of vestibulospinal neurons predominantly reﬂects static encoding of body tilt.

405

The utricle is indispensable for the bulk of vestibulospinal neuron

406

responses

407 Loss-of-function experiments assaying both behavior (Bianco et al., 2012; Ehrlich and Schop408 pik, 2019; Mo et al., 2010) and neuronal responses (Liu et al., 2020) support the proposal
409 that in larval zebraﬁsh, the bulk of the vestibular response is derived from a single vestibular
410 end-organ: the utricle. However, TIPM is inherently multimodal, and might activate other411 systems in addition to the utricle. Angular accelerations can be transduced by the semi412 circular canals. While the semicircular canals are too small to be activated under natural
413 conditions (Beck et al., 2004; Lambert et al., 2008), they can be activated by suﬃciently
414 strong stimuli in comparably small vertebrates (Branoner and Straka, 2014). Translational
415 forces along the body might be encoded by the lateral line (Dijkgraaf, 1963). Finally, pressure
416 along the body might be encoded by the trigeminal system (Ribera and Nüsslein-Volhard,
417 1998). As vestibulospinal neurons are known in other animals to receive a wide variety of
418 multimodal inputs (Sarkisian, 2000) we sought to clarify the role of utricular sensation.
419

We adopted a genetic loss-of-function approach to assay the contribution of the utricle

420 to vestibulospinal responses. Mutants in otogelin, also known as rock solo ﬁsh (Whitﬁeld et
421 al., 1996), do not form a utricular otolith in the ﬁrst 10 days (Roberts et al., 2017). otogelin
422 is selectively expressed in the inner ear (Stooke-Vaughan et al., 2015), avoiding oﬀ-target
423 confounds. We tested the responses of rock solo mutants to both a 20° step and impulse
424 stimulus. We provided both ipsilateral and contralateral impulses; as we previously observed
425 no systematic diﬀerences we aggregated the data to assay responses.
426

We observed that vestibulospinal responses to ipsilateral roll steps in rock solo ﬁsh were

427 severely compromised. The rock solo mutants were less likely to show signiﬁcant changes
428 in ﬂuorescent intensity following a 20° ipsilateral step compared to their wildtype siblings

429 (96% responsive WT, n=24/25 neurons from N=3 ﬁsh; 42% responsive mutants, n=13/31
430 neurons from N=3 ﬁsh). When there were supra-threshold responses, the magnitude of peak
431 ﬂuorescence in mutants was strongly attenuated (2.7±2.3 ΔF/F WT vs 0.21±0.24 ΔF/F
432 mutants, 3-way ANOVA Interaction Eﬀect¡0.001, post-hoc test p=5.9x10-8 ) (Figure 5A,5B).
433

In contrast, responses were not signiﬁcantly diﬀerent between wildtype siblings and rock434 solo mutants following contralateral steps (0.38±0.41 ΔF/F WT vs 0.26±0.30 ΔF/F mu435 tant), nor were responses signiﬁcantly diﬀerent to impulse steps (0.42±0.48 ΔF/F WT vs
436 0.32±0.39 ΔF/F mutant; post-hoc test p=0.99) (Figure 5A,5B) We conclude that contralat437 eral eccentric and impulse responses are predominantly driven by extra-utricular sources.
438 Following both contralateral steps and impulse stimuli, we observed a decrease in the frac439 tion of neurons that responded to the stimulus in rock solo ﬁsh (Contralateral step = 72%
440 responsive WT vs 39% responsive mutant, Impulse = 72% responsive WT vs 44% responsive
441 mutant). Changes to the fraction of neurons that have supra-threshold responses reﬂect an
442 increase in variability of baseline calcium ﬂuctuations in rock solo mutants (baseline SEM:
443 0.019 WT vs 0.032 rock solo), consistent with electrophysiological observations (Hamling et
444 al., 2021; Liu et al., 2020).
445

We conclude that the changes in ﬂuorescence we observe in vestibulospinal neurons fol-

446 lowing ipsilateral body tilts predominantly reﬂects utricular transduction.

447

Vestibulospinal neuron responses develop systematically

448 A distinct advantage of TIPM is its minimally-invasive nature. As such, it is well-suited
449 for experiments that require monitoring the same neurons across multiple timepoints. We
450 asked if TIPM could detect developmental changes in individual vestibulospinal neurons on
451 two diﬀerent days. Prior behavioral work established that larval zebraﬁsh use vestibular
452 information to balance and locomote in diﬀerent ways at 4 and 7 dpf (Ehrlich and Schoppik,453 2017, 2019). We therefore picked 4 and 7 days to assay for diﬀerences in body tilt-evoked
454 responses.

455

We imaged ﬂuorescence after return to horizontal from 10°, 20°, and 30° step stimuli in the

456 same ﬁsh at two ages: 4 and 7 dpf. We were able to reliably identify the same neurons across
457 imaging sessions (Figure 6A). Peak calcium responses within the same neuron were correlated
458 (ρ=0.51) between 4 and 7 dpf (n=71 cells, N=10 ﬁsh)(Figure 6B), but were more variable
459 than neurons in repeated imaging sessions performed on the same day (Figure 6B, gray460 bar) suggesting developmental changes in neuronal encoding. We observed that responses
461 were more variable in our second imaging session (7 dpf, median CV=0.29) than the ﬁrst
462 (4 dpf, median CV=0.19); we therefore chose to compare the slope of peak ﬂuorescence463 responses (“Roll Sensitivity,” a common metric of sensory encoding capacity (Lannou et
464 al., 1979)) of neurons across development, instead of a metric like mutual information that
465 takes response variability into account (Quiroga and Panzeri, 2009). Across all neurons that466 responded to the stimulus at either age, calcium response sensitivity to ipsilateral eccentric
467 rolls strengthened between 4 and 7 dpf (mean slope = 0.07 vs 0.10 ΔF/F/°, Repeated468 Measures ANOVA post-hoc test p=0.002; n=70 cells), and sensitivity to contralateral roll469 did not decrease signiﬁcantly (mean slope = 0.02 vs 0.01 ΔF/F/°, p=0.09). Our data suggest
470 that the population of vestibulospinal neurons improves its ability to encode eccentric roll
471 tilts during this developmental window.
472

Additionally, beyond eﬀects on the population, our longitudinal imaging paradigm al-

473 lowed us to ask 1) how an individual neuron’s sensitivity to ipsilateral or contralateral eccen474 tric roll angle changed across development and 2) whether these changes are systematically
475 patterned. First, we investigated whether developmental changes within the population of
476 vestibulospinal cells was homogeneous. To do so, we examined the distribution of devel477 opmental sensitivity changes in individual cells. We then identiﬁed cells with a signiﬁcant
478 change in roll sensitivity by comparing the observed change in a cell’s sensitivity to a cutoﬀ
479 generated from sensitivity changes in age-shuﬄed data (Methods). We found that individual
480 cells had heterogeneous and asymmetric sensitivity changes to ipsilateral rolls (Figure 6D,
481 top). In response to ipsilateral rolls, individual vestibulospinal cells experienced either no482 change in sensitivity (53/70 cells) or a signiﬁcant increase in sensitivity (14/70 cells) but very
483 rarely experienced a signiﬁcant decrease in ipsilateral sensitivity between 4 and 7 dpf (3/70
484 cells). This asymmetry explains the overall increase in ipsilateral responses observed across
485 the whole population at 7 dpf. In comparison, the distribution of contralateral sensitivity
486 changes was heterogeneous and approximately symmetric (Figure 6D, bottom), with most
487 cells experiencing no signiﬁcant change (51/70 cells) and comparable numbers experiencing
488 a signiﬁcant sensitivity increase (8/70 cells) or decrease (11/70 cells). Together, these ﬁnd489 ings allow us to conclude that the vestibulospinal population is not homogeneous in how tilt
490 responses develop. Speciﬁcally, a majority of cells do not change between 4 and 7 days while
491 small sub-populations either increase or decrease their sensitivity in a directional-dependent
492 manner.
493

We then asked whether the developmental changes an individual cell experiences in the

494 ipsilateral and contralateral directions are correlated; we found that there was no signiﬁcant
495 correlation between sensitivity changes across these two directions (Figure 6E) (ρ=-0.14,
496 p=0.25). Together with previous ﬁndings, these data support the existence of diﬀerent
497 patterns of functional development occurring within the vestibulospinal nucleus that are
498 not coordinated between response directions. While ipsilateral responses strengthen and
499 rarely weaken, the same pattern does not apply to contralateral responses. Additionally,
500 developmental change in one direction does not predict change in the opposing direction.
501 The lack of correlation observed between ipsilateral and contralateral developmental changes
502 suggests that the reﬁnement of these two directions are driven by separate mechanisms.
503

To explore potential mechanisms for systematic changes in response properties across de-

504 velopment, we attempted to predict how ipsilateral and contralateral roll sensitivity within
505 a neuron would change based on the responses we observed at 4 dpf. We asked whether neu506 rons with a signiﬁcant, magnitude-dependent response to roll stimuli early in development
507 (“Early-Tuned”) would selectively strengthen or weaken their responses as they develop.
508 For ipsilateral stimuli, there was no signiﬁcant diﬀerence between sensitivity change distri-509 butions when split between early-tuned (42/70 neurons) and non-tuned cells (Figure 6F,
510 top) (two-sample Kolmogorov-Smirnov test, p=0.99). In contrast, we found that neurons
511 that were early-tuned for contralateral stimuli (24/70 cells) had a signiﬁcantly diﬀerent dis512 tribution of developmental sensitivity changes compared to non-tuned neurons (Figure 6F,
513 bottom)(two-sample Kolmogorov-Smirnov test, p=2.06x10-7 ). Speciﬁcally, early-tuned con514 tralateral neurons were only observed to decrease (11/24 neurons) or have no change (13/24
515 neurons) to their contralateral sensitivity between 4 and 7 dpf, and never strengthened their
516 contralateral responses. Neurons that did not have signiﬁcant contralateral responses at 4517 dpf either strengthened their contralateral sensitivity between 4 and 7 dpf (8/46 neurons) or
518 experienced no change (38/46 neurons). We can therefore conclude that contralateral, but
519 not ipsilateral, sensitivity changes are patterned by a neuron’s responses early in develop520 ment. Such inferences are new to the vestibular ﬁeld and are made possible by our ability to
521 follow the same neurons over multiple days. We propose that our novel approach is there522 fore well-suited for discovering biologically-relevant changes responsible for improvements to
523 neuronal control of posture as animals develop.

524

Discussion

525 Here we report a new method, Tilt In Place Microscopy, to measure neuronal responses
526 following vestibular stimulation. To test TIPM, we mounted a larval zebraﬁsh on a rotat527 ing platform (mirror galvanometer) and measured ﬂuorescence as the ﬁsh returned from an
528 eccentric tilt. Consistent with prior work (Peterson, 1970; Rovainen, 1979), we observed reli529 able responses that vary with tilt magnitude. We tested the reproducibility of our method by
530 mounting the same ﬁsh repeatedly, ﬁnding that TIPM produced little change in the strength
531 and reliability of neuronal responses. By imaging the same neurons both at an eccentric angle
532 and at the horizon, we conﬁrmed that the response reﬂected the steady-state activity while
533 tilted. We next delivered impulse steps, and discovered topographically-organized responses.

534 Consistent with other work, vestibulospinal neurons in mutant zebraﬁsh without utricles re535 sponded minimally following tilts. Finally, we measured activity from a set of vestibular
536 neurons at both 4 and 7 days post-fertilization and reveal systematic changes in sensitivity
537 and selectivity across time. Below we compare TIPM to other approaches/apparatus, discuss
538 limitations and potential extensions, and contextualize our ﬁndings.

539

Comparison to other apparatus/approaches

540 Other groups have tackled the challenge of imaging neural activity while measuring vestibu541 lar responses; each approach requires considerable technical expertise and ﬁnancial resources.
542 One approach involved adapting a powerful technique, the optical trap, to directly displace
543 the utricle (Favre-Bulle et al., 2018). Another approach cleverly miniaturized the hardware
544 so as to permit an entire light sheet microscope to rotate stably (Migault et al., 2018).
545 Together, these apparatus allowed these investigators to characterize vestibular responses
546 across an entire vertebrate brain – a considerable advance. Recently, another group devel547 oped a rotating stage that allowed them to examine the vestibular periphery (Tanimoto et
548 al., 2022). Finally, a complementary approach used for in vivo electrophysiology may be
549 compatible with imaging provided that the microscope was suﬃciently small: mounting the550 preparation on an air-bearing sled (Liu et al., 2020). All four solutions require a familiar551 ity with optical physics and/or engineering to implement and calibrate. All four require
552 specialized and expensive hardware.
553

In contrast, TIPM oﬀers a number of advantages. It is comparatively low-cost and com-

554 patible with any microscope (multiphoton or otherwise) with a long working distance objec555 tive that can accommodate a simple rotating platform. Importantly, the only requirement
556 to control the stimulus is the ability to deliver an analog voltage to control the galvanometer
557 and a digital pulse for calibration. We propose that TIPM’s simplicity, ﬂexibility, and low
558 cost facilitates the study of the neuronal encoding of vestibular responses.

559

Limitations of TIPM

560 The design choices we made facilitate certain experiments, but are not without their own
561 trade-oﬀs. TIPM was designed to image the response to tilts at one particular orientation,
562 to facilitate comparison to baseline measurements so crucial for imaging ﬂuorescence from563 calcium indicators. Responses must be therefore measured after stimulation is complete.
564 Further, we chose to move the preparation relative to a stationary microscope objective. The
565 light path to the same neuron will change with each change in orientation. Consequentially,
566 comparisons across orientations (as in Figure 3) requires normalization to a baseline that
567 accounts for diﬀerent scattering such as an anesthetized baseline (Methods). The inability to
568 measure during stimulation and the challenge of comparing across orientations means that569 our method is likely incompatible with a sinusoidal rotation.
570

The vestibular periphery has been been modeled as an linear, time-invariant system

571 (Laurens et al., 2017). By and large, the measurements underlying this powerful framework
572 are derived from sinusoidal stimulation at diﬀerent frequencies while recording neuronal573 responses across vestibular areas. However, sinusoidal stimulation is not the only way to
574 measure the response of a linear, time-invariant system. Impulse stimuli (e.g. Figure 4A)
575 contain power at a wide range of frequencies. Such “click” stimuli are common in characteriz576 ing auditory responses and the head impulse test is common place during clinical evaluation
577 of semicircular canal function (Halmagyi et al., 2017). Similarly, step stimuli such as we have
578 used here allow evaluation of the DC component (i.e. the steady-state response to gravity
579 at a particular orientation). While TIPM as presented here is incompatible with sinusoidal
580 rotation, we propose that evaluating the responses to impulses (Figure 4) and steps (Figure581 1) as we have here will serve comparably for linear systems analysis of the vestibular system.
582

Unlike other systems that can rotate a full circle, TIPM is constrained to a smaller range

583 of angles due to three factors. First, the galvanometer itself can only rotate 40°. Second, our584 preparation relies on surface tension to keep the water between the sample and the objective.
585 In practice rotations greater than 40° relative to the horizon risk spilling the water. Finally,586 steric considerations limit the achievable rotation. To rotate 90°, the platform would have
587 to be suﬃciently narrow so as to ﬁt entirely within the working distance of the objective (2588 mm) Such a narrow platform would be unwieldy to mount and hold too little water. We
589 therefore do not believe our apparatus will be able to tilt much beyond what we report here.
590 Experiments that require a wider range of angles are better performed on apparatus that591 can rotate more.

592

Ways to extend TIPM

593 For imaging experiments, the choice of indicator and ﬁeld of view set fundamental limita594 tions in time and space. Here we used a slow calcium indicator (GCaMP6s) to measure
595 neuronal activity. All our estimates of vestibular response are convolved with the spike-to596 calcium kernel (Chen et al., 2013). This low-pass ﬁlter constrains our ability to measure
597 vestibular responses regardless of whether stimuli are sinusoidal, impulses, or steps. Recent
598 advances in genetically-encoded voltage indicators suggest that ﬂuorescent imaging of mem599 brane potential is on the horizon (Böhm et al., 2022), or perhaps here (Liu et al., 2022).
600 As TIPM delivers rapid changes to tilt, and is straightforward to integrate with advanced
601 microscopes, we anticipate that it will be ideal for voltage imaging experiments. Similarly,
602 TIPM translates readily to microscopes with wider ﬁelds of view facilitating “whole-brain”
603 approaches.
604

TIPM can accommodate a wide variety of existing hardware to accommodate in vivo

605 imaging in diﬀerent preparations. While we used a mirror galvanometer, any device that
606 can rapidly and precisely rotate away from and back to a given angle will work. Small
607 direct-drive rotation mounts such as Thorlabs DDR25, or larger options such as Newport’s
608 RGV100 series oﬀer rapid and precise rotation and allow for larger loads than the mirror
609 galvanometer. A low-cost option is similarly available by substituting a DC stepper motor
610 and a driver with micro-stepping capability to permit smooth acceleration. Both options
611 would also permit compatibility with current platforms for in vivo imaging in Drosophila

612 (Aragon et al., 2022) and Caenorhabditis (Smith et al., 2022). Naturally, our approach
613 is compatible with head-mounted microscopes (Aharoni and Hoogland, 2019; Zong et al.,614 2022) and stably-mounted high-density probes of electrical activity (Steinmetz et al., 2021)
615 in rodents. We anticipate that labs looking to adopt TIPM will select the device that best
616 rotates their existing preparation.
617

TIPM can be easily extended to permit measuring tail, ﬁn, and/or eye movements in

618 zebraﬁsh. Similar adaptations allow for measurement of leg/wing movements in other ani619 mals. First, it is necessary to replace the mirror galvanometer with a transparent platform.
620 A camera mounted below the apparatus can measure the tail bends and eye movements in
621 the horizontal plane with freely available software such as Stytra (Štih et al., 2019). To
622 measure eye movements in the torsional plane, a glass coverslip can be glued perpendicular
623 to the plane of the slide. A camera can then measure torsional eye movements that follow
624 pitch tilts, as done in (Bianco et al., 2012). As tilt stimuli reliably elicit compensatory pos625 tural and ocular behaviors, such apparatus would provide valuable context to the measures
626 of neuronal activity we report here.

627

Insights into encoding of roll tilt by developing vestibulospinal neu-

628

rons

629 The responses to roll tilts in vestibulospinal neurons reported here largely agree with and ex630 tend prior reports, bolstering conﬁdence in TIPM, and go on to describe novel development
631 ﬁndings that have not previously been observed in vestibular nuclei in any species. We see
632 much stronger responses to ipsilateral than to contralateral steps, replicating ﬁndings from633 larval zebraﬁsh (Hamling et al., 2021; Liu et al., 2020) and other animals (Peterson, 1970;
634 Rovainen, 1979). Mature vestibulospinal neurons are thought to integrate static otolithic
635 information and dynamic information from the semicircular canals (Sarkisian, 2000). We
636 observe minimal impulse responses, consistent with prior reports that larval zebraﬁsh semi-

637 circular canals are too small to transduce angular accelerations (Beck et al., 2004; Lambert
638 et al., 2008), and with our observation that loss of the utricle profoundly decreases the639 responses to tilts. The small impulses we do see are consistent with more recent work in
640 Xenopus that provided considerably larger accelerations to reveal canal-mediated responses
641 (Branoner and Straka, 2014). We conclude that the measurements of neuronal activity in
642 vestibulospinal neurons that we performed here to test our apparatus are likely a reasonable
643 measure of tilt sensitivity.
644

We report systematic changes in neuronal responses from the same vestibulospinal neu-

645 rons measured at two diﬀerent ages. Our choice of age was guided by prior reports showing
646 behavioral diﬀerences in vestibular-mediated locomotion developing between days 4 and 7
647 post-fertilization (Ehrlich and Schoppik, 2017, 2019). While two timepoints are too few to
648 truly deﬁne a developmental trajectory, we observed that across the population, ipsilateral
649 responses strengthened. Such a trend is consistent with the mature tuning for ipsilateral roll
650 reported for vestibulospinal neurons (Peterson, 1970; Rovainen, 1979). By following the same
651 cells over time, TIPM longitudinal imaging also allows us to make new observations about
652 patterns of functional vestibular development. We found that response sensitivity changes
653 do not appear random but instead follow structured patterns which, for some response types,
654 are related to response properties early in development. Our analyses point the way forward
655 to use early response properties to predict how neurons will change across development.

656

Conclusion

657 From birth to death, an animal’s sense of gravity and other accelerations profoundly shapes
658 its physiology (Yates et al., 2013) and journey through the world (Angelaki and Laurens,
659 2020). Perhaps unsurprisingly, studies of the vestibular system have informed nearly every
660 aspect of modern systems-level neuroscience (Goldberg et al., 2012a). Advances in imaging
661 neuronal activity have similarly shaped modern neuroscience. Others have brought imaging
662 and vestibular stimulation together with custom microscopes (Favre-Bulle et al., 2018; Mi-

663 gault et al., 2018; Tanimoto et al., 2022), but adoption requires considerable expertise and664 ﬁnancial resources. Here we describe and validate a novel apparatus/analysis approach we
665 call TIPM to image neuronal responses to body tilts. TIPM is comparatively easy to im666 plement, compatible with a large set of existing microscope designs, low-cost, non-invasive,
667 extensible to a wide variety of preparations, and compatible with longitudinal measurements.
668 We support this claim by conﬁrming and extending our understanding of tilt representation
669 by developing vestibulospinal neurons in the larval zebraﬁsh. Speciﬁcally, we observed pref670 erential sensitivity to tonic stimulation, rough topographic organization, tractable levels of
671 extrinsic variability, and systematic changes across early development. While not without
672 trade-oﬀs, we hope that the simplicity and broad compatibility of TIPM will democratize
673 the study of the brain’s response to destabilization, particularly across development.

Acknowledgments
Research was supported by the National Institute on Deafness and Communication Disordersof the National Institutes of Health under award numbers DC017489 and DC019554, theNational Institute of Neurological Disorders and Stroke under award number NS125280, by
the Leon Levy Foundation, by the Irma T. Hirschl/Monique Weill-Caulier Trust, by theBrain Research Foundation, and by the Dana Foundation. On Elizabeth M. C. Hillman’s
suggestion, Marie R. Greaney was the ﬁrst person (to our knowledge) to mount a larvalzebraﬁsh on a mirror galvanometer. She and David E. Ehrlich performed pilot experiments
used to develop the paradigm described here. The authors would like to thank Jong-Hoon
Nam, Martha Bagnall, and the members of the Schoppik and Nagel labs for their valuable
feedback and discussions.

Author Contributions
Conceptualization: KH and DS, Methodology: KH, YZ, FA and DS, Investigation: KH,
Visualization: KH Writing: KH and DS Editing: DS, Funding Acquisition: KH and DS,
Supervision: DS.

Author Competing Interests
The authors declare no competing interests.

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Table 1: Stimulus properties

Maximum Angular Velocity
Average Angular Velocity
Maximum Angular Acceleration
Average Duration of Step
Average Duration at Eccentric Angle

Unit
deg/msdeg/msdeg/ms2
ms
-

10°
5.3
4.1
18.6
2.2
15 s

20°
6.5
4.9
20.0
3.9
15 s

30°
7.5
5.5
20.2
5.3
15 s

10° Impulse
5.4
3.9
19.8
5.0
0.3 ms15° Impulse
6.1
4.6
21.2
5.6
0.4 ms30° Impulse
6.8
4.2
20.8
13.0
2.5 msFigure 1: Tilt In Place Microscopy (TIPM) produces reliable, directional, andmagnitude-dependent responses following roll tilts. (A) A 4 day post-fertilization (dpf)larval zebraﬁsh mounted in agarose for roll stimuli on a mirror galvanometer. (B) Schematic of
our experimental paradigm. Baseline ﬂuorescence (used for normalization) is measured when
the platform is horizontal. The galvanometer is then stepped and held at an eccentric angle(“Stimulus”) where ﬂuorescence is not recorded, then quickly returned to horizontal whereupon
ﬂuorescent recording begins (“Response”). (C) Voltage-trace from galvanometer during a 10°,20°and 30°step to the left. (D) Inset of feedback voltage from the galvanometer during a step
to 30°. (E) Slices from a two-photon volume of Tg(nefma:GAL4);Tg(UAS:GCaMP6s) ﬁsh.
Dashed yellow overlays indicate pixels that correspond to analyzed vestibulospinal neurons.
Square yellow inset shows close up of a single analyzed cell (scale bar 20 μm) (F) Normalized
ﬂuorescence traces for all trials of one neuron during baseline and response to an ipsilateral 30°
roll. (G) Distribution of coeﬃcients of variation of peak ΔF/F values across 30° step trials forresponsive neurons (n=69 neurons). (H) Mean normalized ﬂuorescence traces for one neuron
during baseline and response to ipsilateral and contralateral roll steps of varying magnitudes(10°, 20°, 30°). (I) Mean peak ΔF/F responses across all responsive neurons for ipsilateraland contralateral rolls of 10°, 20°, and 30° magnitudes. Error bars ± SEM. (J) Distribution of
directionality indices (Methods) across all responsive neurons.

Figure 2: Variability in responses arises predominantly from intrinsic sources(A) Timeline of an experiment of repeated imaging of the same ﬁsh across two
mounts on the galvanometer. (B) Two-photon volumes of vestibulospinal neurons in aTg(nefma:GAL4);Tg(UAS:GCaMP6s) larva during two sequential mounting and imaging experiments. Colored overlays indicate the same neurons located in two separate volumes taken
across the two mounts. (C) Fluorescence traces from two example cells during the baseline period and following an ipsilateral roll during the ﬁrst and second mount for two neurons (orange
and blue traces correspond to colored neuron overlays). (D) Mean peak response during trials
in the ﬁrst mount experiment are strongly correlated with the mean peak response from thesecond mount (ρ=0.83). (E) Peak response on a single experimental trial is strongly correlatedwith peak response on the subsequent trial within the same experiment (ρ=0.79).

Figure 3: Responses after a step back to horizontal from an eccentric angle arestrongly correlated with activity prior to the step. (A) Schematic of stimuli used
to compare responses after return to horizontal (black) to responses at the eccentric angle(magenta). Solid lines indicate stimulus periods where neurons are in focus, dashed lines
indicate stimulus periods where neurons are out of focus. (B) Concatenated traces of mean
normalized ﬂuorescent responses before, during, and after a ipsilateral roll step of 30° for two
example neurons with diﬀering response temporal dynamics (fast-decay on left, slow-decay onright). (C) The mean ﬂuorescence during the last second of the eccentric step is strongly
correlated with the mean ﬂuorescence during the ﬁrst second upon return to horizontal for
all neurons (n=26 neurons) (ρ=0.94). Example fast and slow-decay neurons in (B) identiﬁed
with a star and square symbol. (D) Distribution of calcium response decay slopes during theeccentric step for all neurons. (E) Distribution of times to peak normalized ﬂuorescence during
the eccentric step for all neurons.

Figure 4: Ventral vestibulospinal neurons respond to impulse stimuli in a nondirectional, magnitude-independent manner. (A) Voltage-trace corresponding to feedback from galvanometer during a 10°, 15° and 30° impulse step to the left. (B) Inset of feedback
trace during the impulse step to 30°. (C) Normalized ﬂuorescence traces for all trials of one
neuron during baseline and response to an ipsilateral 30°impulse step. Note the lower scaleon the vertical axis relative to Figures 1-3 (D) Distribution of coeﬃcients of variation of peak
ﬂuorescent intensity across 30° step trials for responsive neurons (n=22 neurons) (E) Mean normalized ﬂuorescence traces for one neuron during baseline and response to an ipsilateral andcontralateral impulse step of varying magnitudes (10, 20, 30°). (F) Mean peak ΔF/F responses
across all responsive neurons for ipsilateral and contralateral impulse steps of 10, 15, and 30°
magnitudes. Error bars ± SEM. (G) Distribution of directionality indices (Methods) acrossall responsive neurons. (H) Spatial location of impulse-responsive (green) or non-responsive(gray) vestibulospinal somata relative to the Mauthner neuron lateral dendrite in μm. (I)Distribution of dorsoventral position of vestibulospinal neuron bodies relative to the Mauthnerlateral dendrite in μm for impulse-responsive (green) and non-responsive (gray) neurons.

Figure 5: The utricle is indispensable for responses to ipsilateral steps, but notcontralateral steps or impulses (A) Example responses (mean ± SD) to 20° step andimpulse stimuli (top row) from vestibulospinal neurons in wildtype and rock solo mutants. (B)Peak ﬂuorescence responses to ipsilateral step, contralateral step, and impulse steps in both
directions in wildtype and rock solo mutants in responsive (green) and non-responsive (gray)neurons.

Figure 6: Longitudinal imaging suggests systematic changes in the complement of
pre-synaptic inputs in developing vestibulospinal neurons (A) Two-photon volumes ofvestibulospinal neurons in a Tg(nefma:GAL4);Tg(UAS:GCaMP6s) larva during longitudinal
imaging experiments at 4 and 7 dpf. Colored overlays indicate the same neurons located in
the volume across the two timepoints. (B) Mean peak calcium response of all neurons at 4
dpf are correlated with their mean peak response at 7 dpf (ρ=0.51), but is more variable than
expected by remounting alone (ﬁt line ± S.D. of residuals from repeated mounting experimentin Figure 2D). (C) Mean peak ΔF/F responses across all responsive neurons for ipsilateraland contralateral rolls of 10, 20, and 30° magnitudes at 4 (black) and 7 dpf (magenta). Error
bars ± SEM. (D) Probability distributions of longitudinal changes in ipsilateral (top) andcontralateral (bottom) calcium response sensitivity (Methods) (n=70 cells). Dashed verticallines represent cut-oﬀs for signiﬁcant sensitivity change. (E) Longitudinal sensitivity changes
in individual neurons across ipsilateral and contralateral roll tilts. (F) Longitudinal sensitivity
changes for ipsilateral (top) and contralateral (bottom) roll tilts split by neurons that havesigniﬁcant magnitude-dependent responses to roll steps in that direction at 4 dpf (“EarlyTuned,” green) versus non-tuned at 4 dpf (“Non-Tuned,” black).