Mutations in SPTBN2 give rise to spinocerebellar ataxia type 5 (SCA5, OMIM# 600224) (**Table 1**), which is characterized by adult-onset and progressive motor incoordination, postural abnormalities and swallowing difficulties. Autopsy tissue from heterozygotic patients with an in-frame deletion at spectrin repeat 3 showed significant loss of Purkinje cells and a thinner molecular layer in the cerebellum. This mutant βIII-spectrin allele also impaired the synaptosomal localization of the glutamate transporter EAAT4 in Purkinje neurons, which may be an underlying cause of the neurodegeneration (Ikeda et al., 2006). Another nonsense recessive allele in SPTBN2 caused developmental cerebellar ataxia and cognitive impairments during childhood. These phenotypes are distinct from SCA5, and this developmental disorder is called spectrin-associated autosomal recessive cerebellar ataxia type 1 (SPARCA1) (Lise et al., 2012; **Table 1**). Nevertheless, how these pathogenic variants give rise to different neurological signatures remains poorly understood. Additional studies are required to reveal the functions, expression profiles and associated proteins of βIII-spectrin in the nervous system; these studies may reveal how these different SPTBN2 variants give rise to distinct disorders.

Several different pathogenic variants in SPTBN4 were recently described (Knierim et al., 2017; Wang et al., 2018a). These patients all have a remarkably similar disease including congenital hypotonia, developmental delay and intellectual disability from early childhood; some patients also have seizures and central deafness (OMIM# 606214) (Wang et al., 2018a; **Table 1**). Wang et al. (2018a) investigated the consequences of these mutations and found that for many, their capacity to interact with AnkG and to be clustered at the AIS was compromised. For other mutations near the C-terminus, the mutant βIV-spectrin protein was unable to bind to phosphoinositides. These properties are essential to stabilize the cytoskeletal architecture of the AIS and to cluster ion channels at AIS and NoR. Future studies to determine the impact of spectrin pathogenic variants on other AIS functions, such as regulating polarity through control of intracellular trafficking, may help determine the cause of pathological phenotypes in these patients.

#### Proteolysis of Spectrins After Injury

Spectrin breakdown products (SBP) have been widely used as biomarkers for central nervous system injury for three main reasons. First, the Ca2+-dependent cysteine protease calpain is activated after CNS injury and cleaves full length αII-spectrin (280 kDa) into 150 and 145 kDa fragments (Pike et al., 2001). These SBP can be isolated from cerebrospinal fluid, allowing clinical evaluation. Second, the quantitative and temporal expression of SBP are highly correlated with the severity of injury and affected regions as evidenced by multiple injury models in rodents, which renders SBP as useful indicators to evaluate the degree of trauma. Finally and most importantly, the breakdown of spectrin-based cytoskeleton faithfully reflects axonal damage, as shown by the disruption of AIS (Schafer et al., 2009) and axon degeneration (Yang et al., 2013). These strengths render SBP as powerful indicators of axon injury and degeneration.

How is the cleavage of αII-spectrin regulated? There are at least two known mechanisms. First, Src-kinase phosphorylates αII-spectrin at Tyr1176 (adjacent to the SH3 domain and cleavage site) which reduces the susceptibility to proteolysis by calpain (Nicolas et al., 2002; Nedrelow et al., 2003). Second, binding between calpain and its endogenous inhibitor

calpastatin maintains homeostatic enzymatic activity. However, after injury, increased intracellular [Ca2+] tips the balance toward activated calpain which degrades the calpastatin. This exacerbates calpain-mediated proteolysis of αII-spectrin (Yang et al., 2013). Transgenic mice that constitutively overexpress human calpastatin (hCAST) in neurons have reduced levels of αII-spectrin SBP and improved motor performance after injury (Schoch et al., 2012). Exogenous delivery of calpastatin into injured neurons also mitigates axon degeneration (Yang et al., 2013). Many calpain inhibitors have now been developed for therapeutic purposes, although improvements in efficacy and safety are still needed [for review see (Ono et al., 2016)]. It is also important to emphasize that αII-spectrin is but one of calpain's many targets and functional deficits in neurons are not due solely to disruption of the spectrin cytoskeleton.

Increased levels of SBP have been reported in many neurological disorders other than CNS injury, including amyotrophic lateral sclerosis (Yamashita et al., 2012), Guillain-Barré syndrome (McGonigal et al., 2010), Parkinson's disease (Diepenbroek et al., 2014) and acute inflammatory autoimmune demyelinating models (Guyton et al., 2010; Das et al., 2013). Inhibition of calpain activity in these studies generally reduced SBP levels. Future studies that examine not only the integrity of axons, but also the structures whose function depends on specialized spectrin-based cytoskeletons (e.g., AIS and NoR), may help to define the pathophysiology of these diseases and injuries. Additionally, it will be important to

#### REFERENCES

determine whether the protective effects of calpain inhibitors is due to their protection of glial or axonal spectrins.

#### CONCLUSION

Since their original description in erythrocytes, our understanding of spectrins has dramatically increased and these important cytoskeletal proteins are recognized as being essential to the functions of diverse cell types in the nervous system. With the advent of advanced microscopy techniques and genetically modified mice, the roles of spectrins are now becoming much clearer. This knowledge will not only help to define the basic function of the nervous system, but it may also suggest therapeutic strategies for nervous system disorders and injuries that include altered or disrupted spectrin cytoskeletons.

#### AUTHOR CONTRIBUTIONS

Both authors wrote the manuscript.

## FUNDING

This study was funded by the Dr. Miriam and Sheldon G. Adelson Medical Research Foundation (Grant Nos. NIH R01 NS044916 and NIH R01 NS069688).

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2019 Liu and Rasband. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Glutamate Imaging Reveals Multiple Sites of Stochastic Release in the CA3 Giant Mossy Fiber Boutons

Sylvain Rama\*, Thomas P. Jensen and Dmitri A. Rusakov\*

UCL Queen Square Institute of Neurology, University College London, London, United Kingdom

One of the most studied central synapses which have provided fundamental insights into cellular mechanisms of neural connectivity is the "giant" excitatory connection between hippocampal mossy fibers (MFs) and CA3 pyramidal cells. Its large presynaptic bouton features multiple release sites and is densely packed with thousands of synaptic vesicles, to sustain a highly facilitating "detonator" transmission. However, whether glutamate release sites at this synapse act independently, in a stochastic manner, or rather synchronously, remains poorly understood. This knowledge is critical for a better understanding of mechanisms underpinning presynaptic plasticity and postsynaptic signal integration rules. Here, we use the optical glutamate sensor SF-iGluSnFR and the intracellular Ca2<sup>+</sup> indicator Cal-590 to monitor spike-evoked glutamate release and presynaptic calcium entry in MF boutons. Multiplexed imaging reveals that distinct sites in individual MF giant boutons release glutamate in a probabilistic fashion, also showing use-dependent short-term facilitation. The present approach provides novel insights into the basic mechanisms of neurotransmitter release at excitatory synapses.

#### Edited by:

Haruyuki Kamiya, Hokkaido University, Japan

#### Reviewed by:

Jon Storm-Mathisen, University of Oslo, Norway Marco Fuenzalida, University of Valparaíso, Chile

#### \*Correspondence:

Sylvain Rama<EMAIL_ADDRESS><EMAIL_ADDRESS>Dmitri A. Rusakov<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Cellular Neurophysiology, a section of the journal Frontiers in Cellular Neuroscience

> Received: 26 February 2019 Accepted: 16 May 2019 Published: 04 June 2019

#### Citation:

Rama S, Jensen TP and Rusakov DA (2019) Glutamate Imaging Reveals Multiple Sites of Stochastic Release in the CA3 Giant Mossy Fiber Boutons. Front. Cell. Neurosci. 13:243. doi: 10.3389/fncel.2019.00243 Keywords: dentate gyrus, CA3 pyramidal cell, short-term plasticity, glutamate release, giant mossy fiber bouton, action potential

## INTRODUCTION

The dentate gyrus is the entry into the hippocampus, with the mossy fibers (axons of granule cells) innervating both CA3 pyramidal cells and stratum-lacunosum interneurons (Acsády et al., 1998). These distinct postsynaptic cell populations are connected through distinct presynaptic elements: "giant" mossy fiber boutons (gMFBs, 4–10 µm) across and their smaller (2–3 µm) variant synapsing onto the thorny excrescences of CA3 pyramidal cells, and relatively small (0.5–2 µm) en-passant boutons and the filopodial extensions emerging from gMFBs both connecting to interneurons (Chicurel and Harris, 1992; Acsády et al., 1998; Rollenhagen et al., 2007). Hippocampal MFB connections show prominent facilitation during repetitive activity and are considered strong "detonating" synapses, generating large postsynaptic responses in CA3 pyramidal cells (Vyleta et al., 2016). This function is sustained by specific morphology, as they show multiple active zones per MFB, and thousands of synaptic vesicles densely packed inside (Acsády et al., 1998; Rollenhagen et al., 2007). They have been widely studied because MF synapses play a key role in processing spatial information such as pattern completion, pattern separation and storage of sequences of events (Kobayashi and Poo, 2004; Bischofberger et al., 2006). Moreover, these synapses provide a key experimental model for analog-digital control of synaptic transmission, linking presynaptic voltage, presynaptic calcium entry, and glutamate release (Geiger and Jonas, 2000;

Alle and Geiger, 2008; Scott et al., 2008). Finally, changes in MF transmission have been a major pathophysiological indicator in epilepsy (Ben-Ari and Represa, 1990; Morimoto et al., 2004) and Down syndrome (Witton et al., 2015). Therefore, an inquisitive exploration of MF synapses helps to understand the fundamentals of synaptic transmission and hippocampal network function in a wide context.

#### RESULTS

### Simultaneous Imaging of Presynaptic Calcium Entry and Glutamate Release at Presynaptic MF Boutons

To directly monitor glutamate release by MFBs, we turned to organotypic slices which we biolistically transfected with the SFiGluSnFR A184V construct (termed iGluSnFR thereafter; see section "Methods"), as detailed earlier (Jensen et al., 2019). With the sparse expression among cells, the iGluSnFR basal fluorescence was sufficient to reveal cell morphology and to track the cell axon. We thus patch-loaded dentate granule cells with the red-shifted calcium-sensitive dye Cal-590 (Tischbirek et al., 2015; Jensen et al., 2019) and followed their axon up to at least 100 µm from the cell soma, toward the CA3 area (**Figures 1A,B**).

In individual cells, we evoked 5 action potentials (APs) at 20 Hz, a bursting pattern similar to that in vivo (Pernía-Andrade and Jonas, 2014; Vyleta et al., 2016; Chamberland et al., 2018). Once we focused on the MF bouton of interest, either of the two imaging protocols was employed: (i) for boutons smaller than 4 µm, a fast "tornado" scanning mode covering the bouton profile (Jensen et al., 2017, 2019), and (ii) for boutons above 4 µm, a straight line-scan along the longest axis of the bouton. In either case, we scanned at 500 Hz and collected both iGluSnFR and Cal-590 fluorescence. Signals were converted as 1F/F values, and the Cal-590 signal was used to confirm AP arrival to the bouton (**Figure 1C**). The iGluSnFR signal revealed glutamate release sites in the bouton, showing release successes and failures (**Figure 1D**). At each individual site, release probability (calculated as the release success rate over all the trials) increased progressively with the AP number in the train, showing classical facilitation (**Figure 1E**). The initial release probability (first AP) at individual release sites was 0.37 ± 0.07 (mean ± SEM, n = 11 boutons), consistent with previous observations (Lawrence et al., 2004).

## Distinct Release Sites Display Stochastic Release

In 6 out of 11 recorded boutons, we could clearly distinguish at least two active release sites, providing rapid glutamate discharges, which did not appear synchronized. We recorded between 3 and 14 consecutive trials (1 min apart) per bouton. Glutamate releases from spatially separate areas were detected in 80 ± 12% trials per bouton (mean ± SEM, a total of n = 43 trials). They were either synchronized or independent (**Figure 2A**). In 25 ± 8% of the recordings (n = 43), we could observe spontaneous (or possibly asynchronous, postburst) release by some release sites (asterisks in **Figure 2** traces). In one gMFB, there were up to four distinct release sites with the first AP inducing glutamate release in only 2 of them and spontaneous release in one of them, independent from the other 3 (**Figure 2B**).

## DISCUSSION

In this brief report, we expressed the SF-iGluSnFR reporter dye in dentate granule cells and monitored glutamate release in gMFBs during brief trains of evoked APs. We detected multiple release sites in individual boutons displaying non-synchronized release activity. This appears in contrast to CA3-CA1 synapses, the majority of which displayed only one detectable glutamate release hotspot (20 out of 23; organotypic slices) whereas the two-hotspot boutons (3 out of 23) showed no detectable asynchrony (Jensen et al., 2019). At the same time, EM data report dual active zones in 38% of CA3-CA1 connections (Shepherd and Harris, 1998), suggesting that super-resolution approaches, such as stochastic localization, once combined with the present technique, might help to better resolve individual release hotspots. Indeed, optical diffraction limit, stereological bias, and limitation of 3D volume scanning should underestimate the occurrence of release sites and therefore their release asynchrony. In some cases, the distinction between en-passant boutons [which are not supposed to have multiple active zones (Acsády et al., 1998)] and the small subtype of giant MF boutons could also be less than clear-cut. Despite of these limitations, the present approach enables us to detect and explore glutamate release and its stochastic features in a direct manner, the task that has hitherto been difficult to achieve.

## Maintenance of High Frequency Transmission by Cross-Talk?

At many synapses, the rates of vesicle fusion exceed the rates of their replenishment leading to vesicle depletion during highfrequency activity (Zucker and Regehr, 2002). However, gMFBs could have formed in a similar way to the Purkinje terminals or the neuro-muscular junction (Telgkamp et al., 2004; Knodel et al., 2014), with large boutons with multiple active zones and densely packed with synaptic vesicles (Rollenhagen and Lübke, 2010). At gMFB synapses, large areas of the opposing pre- and postsynaptic membranes are separated by a narrow cleft. This extracellular space geometry provides, at least in theory, favorable conditions for neurotransmitter spill-over among multiple postsynaptic densities, thus achieving high-fidelity transmission even at low release probabilities and low rates of vesicle replenishment (Vergnano et al., 2014; Zheng et al., 2017). However, our glutamate imaging data suggest that this may not be the case for the MF-CA3 transmission, as the majority of individual glutamate release hotspots in presynaptic boutons do not seem to overlap, at least during first spikes. Indeed, if Purkinje cells do fire at high rates (Telgkamp et al., 2004), it is rarely the case for granule cells of the dentate gyrus (Pernía-Andrade and Jonas, 2014). Moreover, lower release probabilities and readily releasable vesicle pools make hippocampal mossy fiber boutons

en-passant bouton. The fast "Tornado" (spiral) line-scan was set-up to cover most of the bouton visible area. (C) Typical imaging protocol. Traces: Five APs initiated by brief (5 ms) current pulses at 20 Hz (holding voltage −80 mV); Image panels: Tornado line-scans recorded in the Cal-590 (upper) and SF-iGluSnFR (lower) emission channels. (D) Same recordings as in panel (C), but pixel values were averaged over the length of the tornado line-scan and converted as 1F/F. Note that for the Cal-590 signal, each AP induced a calcium entry in the presynaptic bouton (middle) whereas APs 2 and 5 failed to induce a glutamate release (bottom, black arrows). (E) Average release probability (mean ± SEM) for the AP train (n = 11 boutons). Note the increasing probability with the AP number, reflecting presynaptic facilitation.

more suitable for presynaptic facilitation than high-frequency transmission (Delvendahl et al., 2013).

## Implications for Postsynaptic Signal Integration and Presynaptic Machinery

Morphological studies employing 3D electron microscopy have shown that individual giant MF boutons tend to form synaptic connections on more than one postsynaptic CA3 pyramidal cell (Acsády et al., 1998; Rollenhagen and Lübke, 2010). The present finding that individual release sites in such boutons can discharge glutamate relatively independently in a stochastic fashion, suggests that postsynaptic responses in their distinct cell targets could be similarly de-synchronized. It has been previously reported that in juvenile rats, gMFBs may co-release GABA and glutamate (Walker et al., 2001; Ruiz et al., 2003; Beltrán and Gutiérrez, 2012; Münster-Wandowski et al., 2013). The present results thus suggest that GABA and glutamate release from the same gMFB could be, in principle, stochastically separated. Finally, gMFBs show significant structural plasticity in the pilocarpine and kindling models of epilepsy (Danzer et al., 2010; McAuliffe et al., 2011), which could directly affect functional interaction between their individual release sites. What molecular mechanisms underpin such structural and functional changes remains an intriguing and important question.

## METHODS

## Organotypic Cultures of Rat Hippocampus

Hippocampal slice cultures were prepared as described previously (Bialowas et al., 2014). All experiments were carried out according in accordance with the European Commission Directive (86/609/EEC) and the United Kingdom Home Office (Scientific Procedures) Act (1986). Briefly, postnatal day 7–8 Wistar rats were briefly anesthetized by isoflurane inhalation, the brain removed and each hippocampus individually dissected in ice-cold sterile slicing solution consisting (in mM) of Sucrose 105, NaCl 50, KCl 2.5, NaH2PO4 1.25, MgCl2 7, CaCl2 0.5, Ascorbic acid 1.3, Sodium pyruvate 3, NaHCO3 26 and Glucose 10. Hippocampal slices (350 µm) were placed on 20-mm latex membranes (Millicell-CM, Millipore, United Kingdom) inserted

into 35-mm Petri dishes containing 1 ml of culture medium and maintained for up to 30 days in an incubator at 34◦C, 95% O2–5% CO2. The culture medium contained (in ml) 25 minimal essential medium, 12.5 Hanks' balanced saline solution, 12.5 horse serum, 0.5 penicillin/streptomycin, 0.8 glucose (1 M), 0.1 ascorbic acid (1 mg/ml), 0.4 HEPES (1 M), 0.5 B27, and 8.95 sterile H2O. To limit glial proliferation, 5 mM cytosinearabinoside (Ara-C) was added to the culture medium at 4 days in vitro (DIV) for one night.

#### Biolistic Transfection of iGluSnFR Variants

Second generation iGluSnFR variant SF-iGluSnFR.A184V was expressed under a synapsin promoter in dentate gyrus granule cells in organotypic slice cultures using biolistic transfection techniques adapted from manufacturer's instructions. In brief, 6.25 mg of 1.6 micron Gold micro-carriers were coated with 30 µg of SF-iGluSnFR plasmid. Organotypic slice cultures at 8 DIV were shot using the Helios gene-gun system (Bio-Rad) at 120 psi. The slices were then returned to standard culture media the next day and remained for 3–7 days before experiments were carried out.

#### Axon Tracing and Imaging of Pre-synaptic Boutons

We used a Femtonics Femto2D-FLIM imaging system, integrated with patch-clamp electrophysiology (Femtonics, Budapest) and linked on the same light path to two femtosecond pulse lasers MaiTai (SpectraPhysics-Newport) with independent shutter and intensity control. Patch pipettes were prepared with thin walled borosilicate glass capillaries (GC150-TF, Harvard apparatus) with open tip resistances 2.5–3.5 M. Internal solution contained (in mM) 135 potassium methanesulfonate, 10 HEPES, 10 di-Tris-Phosphocreatine, 4 MgCl2, 4 Na2-ATP, 0.4 Na-GTP (pH adjusted to 7.2 using KOH, osmolarity 290–295), and supplemented with Cal-590 (300 µM; AAT Bioquest).

Pre-synaptic imaging was carried out using an adaptation of pre-synaptic glutamate and Ca2<sup>+</sup> imaging methods previously described (Jensen et al., 2017, 2019). Cells were first identified as iGluSnFR expressing using two-photon imaging at 910 nm and patched in whole cell mode as above. Following break-in, 10–15 min were allowed for Cal-590 to equilibrate across the axonal arbor. Axons, identified by their smooth morphology and often tortuous trajectory, were followed in frame scan mode to their targets. Putative Giant Boutons were identified as varicosities on axon collaterals with a minimum diameter and length of ∼2–3 µm (Acsády et al., 1998; Rusakov and Fine, 2003; Rama et al., 2015).

#### REFERENCES

For fast imaging of action-potential mediated iGluSnFR and Cal-590 fluorescence transients at single boutons a spiral shaped ("tornado") scan line was placed over the bouton of interest (described further in the text), which was then scanned at a sampling frequency of ∼500 Hz with excitation at 910 nm. APs were typically induced by trains (20 Hz) of 5 short (5 ms) pulses of depolarizing current (900–1400 pA) in current clamp mode and synchronized with biphoton fluorescent imaging. Ten to fifteen acquisitions were made per boutons, recordings showing APs failures were discarded.

### Data Analysis

After acquisition, iGluSnFr or Cal-590 fluorescence was converted to 1F/F values. Region of Interest (ROIs) were defined by fitting the fluorescence profile with Gaussian equation and extracting the Full Width at Half Maximum (FWHM) of this Gaussian. All of this was calculated by custom-written analysis programs written in Labview (National Instruments) and Matlab (MathWorks).

## DATA AVAILABILITY

All datasets generated for this study are included in the manuscript and/or the supplementary files.

## ETHICS STATEMENT

All experiments were carried out according in accordance with the European Commission Directive (86/609/EEC) and the United Kingdom Home Office (Scientific Procedures) Act (1986).

## AUTHOR CONTRIBUTIONS

SR, TJ, and DR designed the experiments. SR and TJ carried out the patch-clamp and imaging experiments in neurons. SR carried out the analysis. SR and DR wrote the manuscript. All authors contributed to manuscript writing.

## FUNDING

This work was funded by Marie-Curie Fellowship IF-746247 Astromodulation, by the Wellcome Trust Principal Fellowship (212251/Z/18/Z), and by the ERC Proof of Principle grant (767372 – NEUROCLOUD).

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2019 Rama, Jensen and Rusakov. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Dynamic Regulation of Synaptopodin and the Axon Initial Segment in Retinal Ganglion Cells During Postnatal Development

, Dominik Dannehl<sup>1</sup>

,

Annabelle Schlüter1,2, Sabrina Rossberger<sup>3</sup>†

#### Edited by:

Dominique Debanne, INSERM U1072 Neurobiologie des Canaux Ioniques et de la Synapse, France

#### Reviewed by:

Juan José Garrido, Spanish National Research Council (CSIC), Spain Hiroshi Kuba, Nagoya University, Japan

\*Correspondence:

Maren Engelhardt maren.engelhardt@ medma.uni-heidelberg.de

†Present address: Sabrina Rossberger, DELMIC B.V., Delft, Netherlands

#### Specialty section:

This article was submitted to Cellular Neurophysiology, a section of the journal Frontiers in Cellular Neuroscience

> Received: 20 April 2019 Accepted: 28 June 2019 Published: 30 July 2019

#### Citation:

Schlüter A, Rossberger S, Dannehl D, Janssen JM, Vorwald S, Hanne J, Schultz C, Mauceri D and Engelhardt M (2019) Dynamic Regulation of Synaptopodin and the Axon Initial Segment in Retinal Ganglion Cells During Postnatal Development. Front. Cell. Neurosci. 13:318. doi: 10.3389/fncel.2019.00318 Jan Maximilian Janssen<sup>1</sup> , Silke Vorwald<sup>1</sup> , Janina Hanne<sup>4</sup> , Christian Schultz<sup>1</sup> , Daniela Mauceri<sup>2</sup> and Maren Engelhardt<sup>1</sup> \*

1 Institute of Neuroanatomy, Center for Biomedical Research and Medical Technology, Medical Faculty Mannheim, Heidelberg University, Mannheim, Germany, <sup>2</sup> Department of Neurobiology, Interdisciplinary Center for Neurosciences, Heidelberg University, Heidelberg, Germany, <sup>3</sup> Kirchhoff-Institute for Physics, Applied Optics, Heidelberg University, Heidelberg Germany, <sup>4</sup> Abberior Instruments GmbH, Heidelberg, Germany

A key component allowing a neuron to function properly within its dynamic environment is the axon initial segment (AIS), the site of action potential generation. In visual cortex, AIS of pyramidal neurons undergo periods of activity-dependent structural plasticity during development. However, it remains unknown how AIS morphology is organized during development for downstream cells in the visual pathway (retinal ganglion cells; RGCs) and whether AIS retain the ability to dynamically adjust to changes in network state. Here, we investigated the maturation of AIS in RGCs during mouse retinal development, and tested putative activity-dependent mechanisms by applying visual deprivation with a focus on the AIS-specific cisternal organelle (CO), a presumed Ca2+-store. Whole-mount retinae from wildtype and Thy1-GFP transgenic mice were processed for multi-channel immunofluorescence using antibodies against AIS scaffolding proteins ankyrin-G, βIV-spectrin and the CO marker synaptopodin (synpo). Confocal microscopy in combination with morphometrical analysis of AIS length and position as well as synpo cluster size was performed. Data indicated that a subset of RGC AIS contains synpo clusters and that these show significant dynamic regulation in size during development as well as after visual deprivation. Using super resolution microscopy, we addressed the subcellular localization of synpo in RGC axons. Similar to cortical neurons, RGCs show a periodic distribution of AIS scaffolding proteins. A previously reported scaffold-deficient nanodomain correlating with synpo localization is not evident in all RGC AIS. In summary, our work demonstrates a dynamic regulation of both the AIS and synpo in RGCs during retinal development and after visual deprivation, providing first evidence that the AIS and CO in RGCs can undergo structural plasticity in response to changes in network activity.

Keywords: axon initial segment, cisternal organelle, synaptopodin, retinal ganglion cell, visual deprivation

## INTRODUCTION

fncel-13-00318 July 27, 2019 Time: 14:57 # 2

In the retina, visual processing relies on a chain of neurons that transmit information from the outmost photoreceptor layer to the innermost retinal ganglion cell (RGC) layer (Masland, 2012). RGCs convey information via higher order nuclei in the visual pathway to the primary visual cortex (V1) and associated regions. During development, V1 undergoes defined critical periods of cortical plasticity, which shape and refine the mature visual map (reviewed in Hensch, 2004). These critical periods are defined by activity patterns across the visual pathway, and lack of neuronal activity and visual input leads to significant alterations of the final network configuration in V1 (reviewed in Hensch, 2005). While the role of somatodendritic plasticity in this context has been studied extensively (Levelt and Hubener, 2012), it has only recently been acknowledged that axonal plasticity may play an equally important role for the development of sensory cortices (reviewed in Jamann et al., 2017).

The axon initial segment (AIS) is a structurally and molecularly unique axonal domain (Huang and Rasband, 2018; Leterrier, 2018) and is the site of action potential initiation (Kole and Stuart, 2012). Recent work has suggested that it is linked to regulating neuronal excitability on a single cell level, most likely via homeostatic mechanisms (reviewed in Wefelmeyer et al., 2016; Engelhardt et al., 2019). This concept is based on a growing body of evidence from in vitro and in vivo studies and across multiple species showing that the AIS contributes to cellular excitability by increasing or decreasing its length or distance to the soma in response to changes in network activity (reviewed in Yamada and Kuba, 2016; Jamann et al., 2018). These events are thought to be a direct result of changes in synaptic drive either during development or in the mature network: Longer AIS reflect a decrease in synaptic drive, for example after sensory deprivation, and facilitate action potential generation, whereas shorter or distally relocated AIS result from an increase in synaptic drive and are correlated with decreased excitability (reviewed in Jamann et al., 2018).

In work leading to the present study, we described the activitydependent structural maturation of the AIS in V1 pyramidal neurons, which undergo distinct periods of length maturation during the postnatal phase until closure of the critical period of cortical plasticity in mice around P35 (Gutzmann et al., 2014). Visual deprivation led to a significant modulation of this maturation period; in fact, V1 pyramidal neurons were no longer able to reach mature AIS lengths (Gutzmann et al., 2014). The underlying mechanisms that regulate AIS plasticity, in particular length and location changes, remain largely unknown. Recent evidence indicates that intraaxonal Ca2<sup>+</sup> levels may play an important role for AIS plasticity and considering the importance of Ca2<sup>+</sup> currents for the generation and timing of action potentials (Bender and Trussell, 2009), this seems intuitive. Our previous work highlighted a role for the cisternal organelle (CO) in AIS plasticity in V1 pyramidal neurons (Schlüter et al., 2017). The CO consists of stacks of smooth endoplasmic reticulum and is a putative regulator of Ca2+ storage and release in the AIS (Benedeczky et al., 1994; Bas Orth et al., 2007). The structural integrity of the CO is organized by the actin-binding protein synaptopodin (synpo; Bas Orth et al., 2007), which therefore is an excellent molecular marker of this intraaxonal compartment (Sanchez-Ponce et al., 2011; Schlüter et al., 2017).

Currently, it is unclear whether RGC AIS constitute a group of neurons similar to cortical pyramidal cells in terms of AIS morphology, maturation and plasticity, and whether putative AIS plasticity in RGCs could be partially regulated by the dynamic remodeling of the CO. In the present study, we therefore investigated the structural maturation of AIS length and synpo expression in RGCs during retinal development and in dark-reared mice, utilizing a combination of multichannel immunofluorescence, confocal and super resolution microscopy as well as morphometrical analysis. Our data indicate that RGC AIS share a common nanostructure with other excitatory neurons of the visual pathway. Furthermore, RGC AIS develop with similar activity-dependent profiles as shown for V1: Both the AIS and synpo in RGCs undergo periods of dynamic remodeling during retinal development and after visual deprivation, indicating that organelles in the AIS of RGCs can undergo structural plasticity in response to changes in network activity and might therefore impact cellular function during visual processing.

## MATERIALS AND METHODS

#### Animals

The following mouse and rat strains were used: wildtype C57BL/6JRj mice (Janvier Labs, France), B6.Cg-Tg(Thy1- EGFP)MJrs/J mice (own colony at Heidelberg University), and wildtype RjHan:SD rats (Janvier Labs, France). Animals of mixed gender from C57BL/6JRj strain, Tg(Thy1-EGFP)MJrs/J strain, and RjHan:SD strain were maintained with food and water ad libitum on a regular 12 h light/dark cycle. C57BL/6JRj wildtype mice were used for deprivation studies, reared on a 24 h dark cycle as described below, and for the postnatal development study (ages P10 to P > 55). For morphometrical studies of AIS and synpo cluster localization, adult (P > 55) wildtype mice and rats as well as Thy1-EGFP mice were used.

### Developmental Study

For the analysis of AIS length and synpo expression in RGC AIS during retinal development, a total of 6 wildtype mice were analyzed in each of the following age groups: P10, P15, P21, P28, P35, P > 55 (two retinae per mouse, at least 100 AIS per animal <sup>1</sup>= at least 600 AIS/group). For the analysis of AIS in Thy1-EGFP mice, a total of 5 adult (P > 55) mice and a total of 5 control (P > 28) mice were used. For super resolution imaging of the AIS scaffold and synpo clusters in AIS of RGCs, a total of 5 adult (P > 55) wildtype rats and mice were analyzed. Thirty AIS were examined in total. A summary of all experimental groups is given in **Table 1**.

TABLE 1 | Experimental and control groups used in the current study with indication of age of animal, mouse strain, period of visual deprivation, and treatment of tissue for immunofluorescence.

#### Visual Deprivation

Wildtype and Thy1-EGFP mice were kept in completely dark cages with food and water ad libitum. Total absence of light was controlled by exposure of photographic paper located in the cages. Groups of 6 (wildtype) and 11 (Thy1-EGFP) mice each were reared in complete darkness from P0–28 and P0– 35, respectively. Analyses then proceeded immediately after the period of visual deprivation. Data from the developmental study (P28 and 35, wildtype; P > 28, Thy1-EGFP) served as controls for deprivation experiments. All experimental groups are summarized in **Table 1**.

#### Tissue Preparation

Mice were exsanguinated with 0.9% saline under deep anesthesia with ketamin (120 mg/kg BW)/xylazine (16 mg/kg BW). Animals were then perfused transcardially with ice-cold 4% paraformaldehyde (PFA) in 1x PBS. Whole retinas were extracted and processed as follows: Eyes were enucleated with fine curved forceps and were transferred into a 5 cm culture dish containing ice-cold 1x PBS. Retinae were dissected under a stereomicroscope with an internal light source. A hole was cut into the eye at the corneal limbus by using a pair of fine spring scissors. After a circumferential incision along the limbus, the cornea, iris, vitreous body, and lens were removed in toto. The retina was extracted from the eye cup. Retinae were fixed in 4% PFA for 10 min and were washed three times in 1x PBS before further processing.

#### Immunofluorescence

Double and triple immunofluorescence was performed on freefloating whole retinae as described previously (Schlüter et al., 2019). Retinae were blocked for 4-5 h (in 0.5% Triton X-100, 0.2% bovine serum albumin and 0.02% sodium azide in 1x PBS) at 4◦C. Primary antibodies were diluted in dilution buffer (1% Triton X-100, 10% fetal calf serum and 0.02% sodium azide in 1x PBS) according to previously validated concentrations (Gutzmann et al., 2014; Schlüter et al., 2017). Retinae were incubated with primary antibodies for 24 h at 4◦C. Afterward, tissue was washed three times in 1x PBS for 15 min each. Retinae were then incubated with fluorophore-conjugated secondary antibodies diluted in dilution buffer for 24 h at 4◦C, followed by additional washing steps (3x in 1x PBS, 15 min each). Prior to mounting, retinae were fixed for 10 min in 4% PFA and washed in 1x PBS (2x for 5 min). Tissue was then cut from the rim to 1/3 of the radial length and was flat-mounted on slides. All performed stainings were accompanied by negative controls, in which omitting the primary antibody completely abolished all stainings. For confocal and STED microscopy, retinae were embedded in Roti <sup>R</sup> -Mount FluorCare mounting medium or Mowiol (Carl Roth, Karlsruhe, Germany). For SIM and SMLM, retinae were embedded in ProLong <sup>R</sup> Gold mounting medium (Thermo Fisher Scientific, Waltham, MA, United States) or switching buffer [10% dd H2O, 10% 1x PBS, 80% glycerol, 0.1M cysteamine, 100u glucose oxidase (Type VII from Aspergillus niger), 800u catalase (from Bovine liver), 0.1M D-(+)-Glucose]. For shift corrections (SIM and SMLM), 1 µl of FluoSpheres <sup>R</sup> (FluoSpheres <sup>R</sup> Fluorescent Color Kit, Carboxylate-Modified Microspheres, 0.04 µm, four colors; Thermo Fisher Scientific, Rockford, IL, United States) was transferred near the edge of the slide. Coverslips were sealed onto slides using transparent nail polish. All antibodies used in this study are summarized in **Supplementary Table 1**.

#### Confocal Microscopy

Conventional laser scanning confocal microscopy was carried out using a C2 confocal microscope (Nikon, Alzenau, Germany; laser lines: 642, 543, and 488 nm), with a 60x objective (oil immersion, NA 1.4) and a SP5 confocal microscope Leica, Mannheim, Germany; Laser lines: 633, 561, 514, and 488 nm) with a 63x objective (oil immersion, NA 1.4), respectively. To increase the number of in-focus immunoreactive structures, stacks of images were merged into a maximum intensity projection and saved as jpeg and tif format. Thickness of single optical sections was 0.2 µm in stacks of 3–5 µm total depth. Confocal x-y-resolution was constantly kept at 0.21 µm per pixel. Images for qualitative analysis were evaluated and enhanced for contrast in Fiji (Image J) and Photoshop C5 (Adobe Systems, United States).

#### Super Resolution Microscopy

Three different super resolution methods were applied to obtain a comprehensive understanding of the RGC AIS nanostructure.

Stimulated Emission Depletion (STED) imaging was performed using a STEDYCON (Abberior Instruments GmbH, Göttingen) with excitation lasers at 488, 561, and 640 nm, and a STED laser at 775 nm (maximum intensity 1.25 W; all lasers are pulsed with 40 MHz repetition rate). The STEDYCON was mounted on the camera port of an AxioImager.Z2 upright microscope (Zeiss, Jena, Germany), equipped with a 100x objective (alpha Plan-Apochromat, Oil, DIC, Vis, NA 1.46; Zeiss). The pinhole was set to 1.1 Airy units for 650 nm emission. Fluorescence was detected on avalanche photo diodes, with emission bands between 650–700 nm, 578–627 nm, and 505–545 nm, respectively. Data was stored in .obf format and exported as tif files for further analysis.

Correlative Structured Illumination Microscopy (SIM) and Single Molecule Localization Microscopy (SMLM) were performed using a custom-built microscope, which is described in detail in Rossberger et al. (2013). For acquisition, a high

numerical objective (Leica HCX PL APO 100x/1.4 oil CS), a charge-coupled device-camera (Sensicam QE, PCO, Kelheim, Germany) and the following laser lines were used: 568 nm laser line (Coherent Sapphire 568 HP, 200 mW, Coherent, Dieburg, Germany) and 671 nm (VA-I-300-671, 300 mW, Beijing Viasho Technoloy Co. Ltd., Beijing, China). For each laser line, excitation and emission light were separated using an appropriate dichroic mirror (Di02-R568 and Q680LP, both Semrock, Rochester, NY, United States) and emission filter combination (BLP01-561, Semrock and LP XF 3104, Omega Optical, Olching, Germany).

Prior to SIM and SMLM acquisition, sections were scanned for correlated AIS-(568 nm) and Synpo-(647 nm) signal using the widefield mode of the microscope. For this purpose, the 568 nm excitation laser line was used in combination with an edge basic longpass emission-filter (BLP01-568R-25, Semrock, Rochester, NY, United States). The resulting widefield image (pixel size of 64.5 nm) shows an overlay of both color channels within one image and was used for shift corrections of the two color channels recorded separately in super resolution mode. Details of SIM and SMLM image acquisition are outlined in the **Supplementary Methods**.

#### Morphometrical Analysis

We classified RGCs in Thy1-GFP mice where the entire cell could be visualized and measured soma size, dendritic tree diameter and stratification of dendrites into the inner plexiform layer (IPL) according to previously published guidelines (Sun et al., 2002b). We used parameters of soma and dendritic field size to classify RGCs into the different classes (A1–A2, B1– B4, C1–C6, D1–D2; O'Brien et al., 2014) as well as dendritic stratification into the IPL to further isolate A RGCs (RGCA) into ON- and OFF-ganglion cells (Anderson et al., 2016; Saha et al., 2016). Confocal z-stack images of entire RGCs were projected using AutoQuant X3 software (Media Cybernetics). The obtained three-dimensional images were turned 90 degrees to visualize the stratification of the dendritic tree of RGCs within the IPL (**Supplementary Figure S1A**). ON-ganglion cells were identified based on the ramification of their dendritic processes in sublamina b of the IPL whereas OFF-ganglion cells ramify in sublamina a of the IPL. In order to confine our data analysis to the RGC layer specifically and avoid any potential cross-contamination with cells or AIS from other retinal layers, confocal stacks were acquired exclusively within the ganglion cell layer (GCL) of the retina. The GCL could be identified clearly by NeuN immunostaining, nuclei labeling (TOPRO), and its location within 5–10 µm underneath the nerve fiber layer of the mouse retina. Thus, AIS of other retinal layers, such as AIS-like processes from AII amacrine cells located in the inner nuclear layer (Wu et al., 2011) were not included in our analysis. AIS in the GCL were selected only after clear identification of proximal and distal endpoints of immunoreaction to ankyrin-G (ankG) or βIV-spectrin could be traced across optical sections of a stack without any discontinuation or broad stratification into other layers, based on the 3D-projections created for the RGC classification (**Supplementary Figure S1B**). Overlapping AIS were not included in the analysis to avoid any potential interference of immunofluorescent signal with length and distance analysis.

AIS length and distance from the soma were obtained using a self-written program in Python (AISuite et al., unpublished; Hoefflin et al., 2017; Ernst et al., 2018). Briefly, in this application, the AIS is traced manually, beginning with the axon hillock and ending with the axon past visible AIS staining to exclude personal bias. The cut-off threshold for AIS length identification was set to 30% of the individual maximum intensity. Triplets of AIS signal values were analyzed to reduce the influence of upward outliers. The proximal and distal borders of the AIS were determined as the first, respectively the last triplet of signal values which were higher than the defined cut-off. The pixel difference between the proximal and distal end of the AIS was calculated and converted into length in µm taking the microscope's calibration into account. AIS distance to the soma was determined by calculating the distance between the onset of the individual region of interest at the axon hillock and the estimated beginning of the AIS, using the same cut-off. All data were exported as HDF5 and Excel files for statistical analysis.

Periodicity of the AIS scaffold was analyzed similar to previously published methods (D'Este et al., 2015). Briefly, brightness and contrast were adjusted linearly for each STED and SMLM image. Fluorescence intensity plot profiles were measured using ImageJ along a 5 pixel wide line drawn along the AIS. For each AIS, a minimum of 3 regions were measured. The distances between interpeaks were determined using the peak analyze function in OriginPro software (Additive Friedrichsdorf, Germany).

#### Synpo Expression Analysis

Number and size of synpo clusters were measured using a self-written macro in ImageJ as previously published (Schlüter et al., 2017). AIS containing synpo-positive immunofluorescence signals were manually outlined. Background signals beyond the AIS were eliminated. For cluster analysis, the "Color Threshold" option was used. Threshold level was set to 55 and minimum size of pixels was 5. Both values were kept constant during measurements. Synpo-positive clusters per AIS were defined by these parameters and were measured automatically for mean number and size (in pixels). For cluster sizes, mean pixel values were translated into µm<sup>2</sup> (area [µm<sup>2</sup> ] = area [pixels] × (microscope resolution)<sup>2</sup> ).

#### Image Analysis

For analysis of SIM data, nine images of conventional resolution for each z layer and each color channel were recorded. These images were reconstructed and deconvolved using custom developed software written in Matlab (Best et al., 2011). Image reconstruction resulted in one image for each layer within the z-stack and each color channel. For reconstruction of synpo clusters within RGC AIS, 3D SIM images were processed first by blind iterative deconvolution (theoretical PSF based on the optical properties of the microscope and the sample, 10x iteration) according to standard procedures in AutoQuant X3 (Media Cybernetics). Subsequently, to visualize x-y-z information about synpo cluster localization within the AIS,

deconvolved files were reconstructed (surface) using Imaris 9.0 (Bitplane, Zurich) according to previously published protocols (Schlüter et al., 2017).

For analysis of SMLM data, single molecule positions of optically isolated molecules were determined by using custom software written in Python. Briefly, a Gaussian function was fitted to each single molecule signal and its center of mass was calculated. Center coordinates evaluated from all images of the sequence were transferred into one image, marking the position of one fluorophore. The mean position or localization accuracy was 1x = 10.6 nm, resulting in a position image. Images were rendered using a triangulation algorithm as previously published (Baddeley et al., 2010). The area spanned by three next neighbor fluorophore positions (triangle) was translated inversely into intensity values depending on the size of the area. High intensity values were translated into small distances between fluorophore positions and vice versa. Pixel size for visualization was either 5, 32.25, or 64.5 nm. Single molecule positions were randomly jittered to generate 100 slightly different images, which were overlaid afterward to smooth structure edges. Thus, single molecule positions within the images were transferred into an image showing structures in a more conventional sense.

#### Statistical Analysis

Mean values of AIS length and position as well as size and number of synpo clusters per AIS were calculated in Excel (Microsoft). Standard deviation (SD) were calculated and mean values were plotted and analyzed in GraphPad Prism 7 software (GraphPad Software, San Diego, CA, United States). Values were tested for normal distribution by performing Shapiro–Wilk normality test (for testing mean values, n = 6) or D'Agostino and Pearson normality test (for testing single values, n > 600). Unpaired t-test or Mann–Whitney t-test was carried out for statistical comparison of two groups. One-way ANOVA, Kruskal–Wallis or two-way ANOVA test was applied for comparing three or more groups. Post hoc correction was performed by Tukeys or Sidak post test. Boxes extend from the 25th to 75th percentile. Error bars are drawn down to the minimum and up to the maximum value. Asterisks indicate significant differences (∗p ≤ 0.05). For frequency distribution, bin centers were classified by dividing the entire range of values into a series of intervals. Values were then counted and assigned to each interval. For frequency distribution of ankG/β-spectrin periodicity, the entire range was 0–0.4 µm. Bin center steps were 0.02 µm. For frequency distribution of AIS length, the entire range was 10–55 µm. Bin center steps were 2 µm.

#### RESULTS

#### AIS of Mature RGCs Are Characterized by a Distal Localization and a Defined Nanoscale Scaffold

The morphology of RGC AIS in adult mouse retinae was characterized by immunofluorescence against typical AIS scaffolding proteins such as ankyrin-G (ankG) and βIVspectrin, followed by confocal and super resolution microscopic qualitative analysis (**Figures 1**, **2**). We found that both scaffolding proteins are expressed abundantly across all unmyelinated fiber tracts traversing the retina toward the optic disk and colocalize with neurofilament 200 kDa (**Figure 1A**). Individual RGC AIS are clearly distinguishable from the thicker fiber tracts based on their thin structure, increased ankG/βIV-spectrin expression, and decreased neurofilament 200kDa-immunoreaction (**Figure 1A**, magenta arrows). Using the transgenic Thy1 M-GFP mouse line, which sparsely labels RGCs under control of the Thy1 promoter in the retina (Feng et al., 2000), we observed a significant distance between the soma and proximal beginning of the AIS in most RGCs (20 ± 7.1 µm SD; **Figures 1B,D**). This highlights a significant difference to pyramidal neurons in V1, which in mice almost never showed a gap between proximal AIS and soma, but if a gap was detectable, the distance was significantly smaller (P28 visual cortex 2.3 ± 1.5 µm SD; **Figures 1C,D**).

To investigate the nanoscale architecture of the AIS in RGCs, we utilized STED microscopy and Single Molecule Localization Microscopy (SMLM). Analysis revealed that the ankG and βIV-spectrin scaffold is periodically spaced with distances of approximately 181 ± 26.08 nm SD between single fluorescent signal peaks (STED, **Figures 2A–D**; For SMLM: 205.6 nm ± 42.22 nm SD **Supplementary Figures S2A,B,C**). These findings confirm that RGC AIS are of similar nanostructure as other neuron populations, for which a scaffold with ∼190 nm spacing has been reported (D'Este et al., 2015; Leterrier et al., 2015). STED microscopy also revealed a striking deviation in AIS architecture compared to other neuron populations. RGC AIS often appear 'looped' or arranged in a corkscrew manner almost reminiscent of a double-helix structure (**Figures 2A,B**), with large 'gaps' between two single strands of axonal scaffold (**Figure 2A**).

#### AIS of RGCs Undergo Dynamic Length Maturation During Postnatal Development

The structural maturation of RGC AIS during the postnatal retinal development was investigated by immunofluorescence and confocal microscopy combined with AIS length analysis as previously published (Gutzmann et al., 2014; Schlüter et al., 2017). AIS length development was analyzed from P10 until adulthood (P > 55; **Figure 3**). AIS were the longest during the early postnatal period until eye-opening around P15 (P10: 24.45 ± 0.42 µm SD, P15: 24.11 ± 0.28 µm SD; **Figures 3A,D**). From P21 onward, AIS length significantly decreased, reaching stable length from P28 until adulthood (P21: 20.68 ± 0.36 µm SD, P28: 16.92 ± 0.28 µm SD, P > 55: 16.82 ± 0.25 µm SD; **Figures 3B,D**). In parallel, the cumulative length distribution of retinal AIS significantly shifted from a heterogeneous distribution to a homogenous length distribution at P21 (**Supplementary Figures S3A–H**). Across the three adult stages analyzed, RGC AIS length averaged at 16.9 µm ± 0.6 µm SD (P28, P35, and P > 55; **Figure 3D**), indicating that RGC AIS are shorter than AIS of other cell populations in adult wildtype

Insert: magnification of boxed region in (B), indicating the gap between proximal AIS onset and the soma (upper panel, white arrows) and the length of the entire AIS (lower panel, magenta arrows). (C) Image of V1 cortical pyramidal neurons from adult mouse stained for NeuN (green) and βIV-spectrin (magenta), with only a small distance between soma and proximal AIS indicated by white arrows. (D) Quantification of the soma to AIS distance from V1 cortical neurons and RGCs. Unpaired t-test. <sup>∗</sup>p ≤ 0.05, n = 5 animals (at least 100 AIS per animal). Scale bar in (A) = 20 µm, (A1) = 10 µm, (B) = 30 µm, (B1) = 10 µm, (C) = 20 µm.

mice, such as the visual cortex (average approximately 33 µm; Gutzmann et al., 2014), hippocampus (average approximately 29 µm; Kaphzan et al., 2011), or substantia nigra (average approximately 26 µm; Gonzalez-Cabrera et al., 2017).

## Synaptopodin Is Expressed in AIS of RGCs During Retinal Development

A subset of AIS of hippocampal and cortical neurons express the Ca2+-storing CO (Bas Orth et al., 2007; Sanchez-Ponce et al., 2012a; Schlüter et al., 2017). Considering the difference in morphology and function between RGCs and upstream neurons of the visual pathway, we asked whether RGC AIS also contain such intraaxonal Ca2+-stores. We applied immunofluorescence in retinal whole mounts to test whether synaptopodin (synpo), a core structural component of the CO, is expressed in RGC AIS (**Figure 4**). Interestingly, we found robust synpo expression in a subset of AIS of Thy1-positive RGCs and always confined to the borders of the AIS (**Figures 4A–D**).

A previous study using super resolution microscopy in cortical neurons proposed that synpo-positive COs are clustered at ankG-deficient sites in the AIS (King et al., 2014). We addressed the question whether these clusters appear at sites of ankG-deficiency in RGC AIS. Indeed, occasionally we found synpo clusters in the vicinity of gaps in the axonal scaffold (**Figures 4B,C** and **Supplementary Figure S4B**). However, by applying super resolution microscopy, we found that synpopositive clusters are often located between rings of βIV-spectrin within the AIS (**Supplementary Figure S4A**). Occasionally, synpo clusters appeared as if they were located outside of the AIS (**Figure 4C**, arrowhead).

## The Size, but Not the Number of Synpo Clusters in AIS of RGCs Undergoes Changes During Retinal Development

In visual cortex neurons, synpo clusters undergo a dynamic regulation in both size and number during development (Schlüter et al., 2017). Here, we investigated synpo expression in RGC AIS during postnatal retinal development. Synpo expression in AIS was first observed at P10 in 26.5 ± 6.54% SD of RGC AIS (**Figures 5A,C**). During further postnatal development, the percentage of synpo-positive AIS in the retina remained stable and did not undergo any significant changes (i.e., P28: 28.17 ± 5.78% SD; **Figures 5B,C**). These data suggest that the presence of synpo in RGC AIS is not influenced by visual input mediated by the eye-opening phase around P13,14.

However, it remained an open question whether synpo clusters undergo similar dynamic changes during development as in V1 (Schlüter et al., 2017). Therefore, we examined the developmental changes of synpo expression in RGC AIS and quantified the number (**Figures 5D,E**) and size (**Figures 5F,G**) of synpo-positive clusters per AIS. At P10, the average number of synpo clusters per AIS was 1.6 ± 0.05 SD (**Figure 5E**). During further retinal development, the number of synpo clusters remained stable (i.e., P35: 1.53 ± 0.03 SD; **Figure 5E**), recapitulating the finding regarding the percentage of synpoexpressing AIS during retinal development (**Figure 5C**). The size of synpo-positive clusters in RGC AIS was 0.53 ± 0.01 µm<sup>2</sup> SD at P10 and remained stable until P21 (0.59 ± 0.01 µm<sup>2</sup> SD; **Figure 5G**). In comparison to the stable development of number of synpo-positive clusters per AIS, the size significantly decreased from P21 to P28 and P35, respectively (P28: 0.50 ± 0.02 µm<sup>2</sup> SD, P35: 0.52 ± 0.004 µm<sup>2</sup> SD; **Figure 5G**). During further postnatal development, synpo cluster sizes reached a plateau, which was maintained throughout adulthood (P > 55: 0.53 ± 0.02 µm<sup>2</sup> SD; **Figure 5G**).

### Synpo Expression in RGC AIS Seems to Reduce and Stabilize AIS Length During Retinal Development

Since synpo-positive AIS display shorter AIS length and reduced dynamic length maturation during visual cortex development (Schlüter et al., 2017), we analyzed the developmental changes of length in the subset of synpo-expressing RGC AIS (**Figure 6**), and compared it to the length maturation of the entire

AIS population in the retina (**Figure 3**). At P10, synpopositive AIS have an average length of 15.48 ± 0.35 µm SD (**Figures 6A,C**). During further postnatal development, AIS length continuously decreased (i.e., P28: 14.60 ± 0.25 µm SD; **Figures 6B,C**) and reached a plateau in adulthood (P > 55: 13.91 ± 0.12 µm SD; **Figure 6C**), at which time AIS length is significantly reduced compared to the early postnatal period at P10. In parallel, the cumulative length distribution of

bar (A) = 30 µm, (B,C) = 5 µm.

ANOVA. <sup>∗</sup>p ≤ 0.05, n = 6 animals (at least 100 AIS per animal). Scale bar (A,B) = 20 µm, (C,D) = 5 µm.

RGC AIS was significantly homogeneous already early in development (**Supplementary Figures S5A–H**). Further, the average length of synpo-positive AIS was significantly shorter during retinal development as compared to the entire AIS population [**Figure 6C**, compare to **Figure 3D**; i.e., P21 (all AIS) vs. P21 (synpo<sup>+</sup> AIS: 20.68 ± 0.88 µm SD vs. 14.75 ± 1.02 µm SD)]. These findings suggest that similar to the AIS of pyramidal neurons in the visual cortex (Schlüter et al., 2017), the presence of synpo-positive clusters within the AIS might reduce and possibly stabilize AIS length in RGCs during postnatal retinal development.

### Visual Deprivation During Retinal Development Impairs AIS Length Maturation in the Retina

Recent studies have demonstrated that sensory deprivation prevents the structural maturation of the AIS in auditory, visual

and somatosensory system neurons during development (Kuba et al., 2010; Gutzmann et al., 2014; Schlüter et al., 2017; Jamann and Engelhardt, unpublished). In fact, AIS maturation seems to be an activity-dependent process during which network activity contributes to the final mature length of AIS in cortical principal neurons (reviewed in Jamann et al., 2018). Considering the significant AIS length reduction we observed after eye-opening around P15 in RGC AIS (**Figure 3**), we hypothesized that a similar effect also contributes to AIS length maturation in the retina. To test this, we used visual deprivation protocols and reared mice in complete darkness from birth until P28 and P35, respectively. We measured AIS length in these animals and compared it to that of control animals kept under normal light/dark conditions (**Figure 7**). Similar to the observations during visual cortex development (Gutzmann et al., 2014), darkrearing led to a significant increase in AIS length at P28 and P35 (P28 control vs. dark: 16.90 ± 0.28 µm SD vs. 25.69 ± 0.39 µm SD, **Figures 7A–C**; P35 control vs. dark: 16.92 ± 0.18 µm SD vs. 25.62 ± 0.31 µm SD, **Figure 7C**).

Of note, RGCs are heterogeneous regarding their morphology, firing properties and function. We therefore used data obtained from Thy1-GFP mice to further classify the different RGCs in our dataset as outlined in the methods. We found that most Thy1 positive RGCs belong to class A1 and A2, whereas other classes were represented only by up to ca. 10% of cells (**Supplementary Figure S1C**). Due to the low number of RGCB, RGCC, and RGC<sup>D</sup> cells, sufficient detection of reliable numbers of cells and related AIS in these classes was not pursued.

Selecting only RGC<sup>A</sup> neurons for analysis, we found similar AIS elongation after dark rearing in this subtype [**Figures 7D– F**; control (P > 28) vs. dark (P28): 24.29 ± 1.95 µm SD vs. 28.33 ± 2.22 µm SD]. In these same cells, no significant difference in the gap between proximal AIS and soma was observed [**Supplementary Table 2**; control (P > 28) vs. dark (P28): 22.70 ± 3.44 µm SD vs. 22.80 ± 4.06 µm SD]. Size frequency histograms of AIS length in all RGC classes further highlighted that dark-rearing until P28 and P35 led to an AIS length distribution similar to that found in young animals, suggesting that neurons maintain a juvenile AIS length (**Supplementary Figures S3A–H**). Thus, visual deprivation seems to prevent the structural maturation and developmental shortening of AIS length in RGCs during postnatal periods.

## Visual Deprivation During Retinal Development Increases Synpo Expression and Length of Synpo-Positive RGC AIS

We next hypothesized that visual input influences not only AIS length maturation, but also the development of synpo clusters in AIS of RGCs. Thus, we compared changes in synpo protein expression in RGC AIS in control animals with animals subjected to dark-rearing for 28 and 35 days (**Figures 8A–G**). We quantified the percentage of synpo-expressing AIS as well as the number and size of synpo-positive clusters per AIS. Regarding the percentage of synpo-expressing AIS, we found that the subset of AIS that contain synpo-positive clusters increased after dark-rearing for 28 and 35 days, which was significant in P35 animals (P28 control vs. dark: 28.17 ± 2.36% SD vs. 36.83 ± 3.50% SD, **Figure 8D**; P35 control vs. dark: 24.0 ± 2.41% SD vs. 32.8 ± 1.2% SD; **Figure 8D**). Further, dark-rearing for 28 days resulted in a significant increase in the average number of synpo clusters per AIS (P28 control vs. dark: 1.53 ± 0.03 SD vs. 1.71 ± 0.04 SD; **Figures 8A,B,E**) as well as significant increase of synpo cluster size per AIS (P28 control vs. dark: 0.50 ± 0.02 µm<sup>2</sup> SD vs. 0.58 ± 0.01 µm<sup>2</sup> SD; **Figures 8A,C,F**). Longer periods of visual deprivation until P35 resulted in an unchanged number of synpo-positive clusters per AIS as compared to the control conditions (P35 control vs. dark: 1.53 ± 0.03 SD vs. 1.46 ± 0.07 SD; **Figure 8E**). Instead, an increase of the size of synpo-positive clusters in AIS of P35 darkreared mice was found when comparing them to P35 control mice and visually deprived P28 mice, respectively (P35 control vs. dark: 0.53 ± 0.004 µm<sup>2</sup> SD vs. 0.65 ± 0.04 µm<sup>2</sup> SD, **Figure 8F**; P28 dark: 0.58 ± 0.01 µm<sup>2</sup> ; **Figure 8F**). These data suggest that visual

input can influence number and size of synpo-positive clusters in AIS of RGCs during postnatal development. Moreover, synpo clusters might fuse within the AIS by further increasing in size when visual input is lacking for periods longer than P28 days. Interestingly, sensory deprivation led to an increase in RGCs that express synpo in their AIS; in other words, RGCs that normally would not express synpo, seemed to initiate expression after the lack of visual input during postnatal development.

Our findings indicate a potential AIS length-stabilizing effect of synpo in RGCs during postnatal development (**Figure 6**). Consequently, we questioned whether synpo-positive AIS still retained their ability to adapt their length to changes in visual input as seen for the entire retinal AIS population (**Figure 7**). Therefore, we quantified the length of AIS in the subset of synpo-expressing RGCs in mice that were dark-reared for 28 and 35 days and compared them to control mice raised under normal light/dark conditions (**Figure 8G**). Under all dark-rearing conditions, a significant increase in the length of synpo-positive AIS was observed (P28 control vs. dark: 14.60 ± 0.25 µm SD vs. 16.96 ± 0.28 µm SD, **Figure 8G**; P35 control vs. dark: 14.16 ± 0.27 µm SD vs. 15.68 ± 0.40 µm SD; **Figure 8G**). Further, length distribution histograms of synpo-positive AIS highlighted that visual deprivation led to a significant shift from homogenous to heterogeneous length distributions (**Supplementary Figures 5A–H**). In conclusion, even though the presence of synpo in AIS of RGCs seems to stabilize AIS length during postnatal development under physiological conditions, synpo-positive AIS maintain the ability to dynamically adjustment their length after loss of visual input.

#### DISCUSSION

In V1, where visual input is processed, the AIS of pyramidal neurons is highly plastic during development and after sensory deprivation (Gutzmann et al., 2014). For RGCs, which constitute a downstream neuron population in the visual pathway, the precise morphology as well as putative plasticity of AIS has not been studied in detail so far. In the present work, we investigated the structural maturation of AIS and the CO in

RGCs during retinal development and after sensory deprivation. Our work highlights that (1) RGC AIS have a periodically spaced scaffold of ankG/βIV-spectrin rings; (2) AIS length in RGCs changes with the maturation of the retina during postnatal development; (3) RGC AIS express the specific CO marker synpo, which is located within the ankG/βIV-spectrin scaffold occasionally in close proximity to ankG/βIV-spectrin deficient sites; (4) the percentage of synpo-positive AIS is stable during development, but synpo cluster sizes change with AIS maturation; and (5) visual deprivation prevents the maturation of AIS shortening in RGCs and increases synpo expression in AIS during postnatal development.

Taken together, our data indicate that the presence of synpo could provide structural stability to RGC AIS during periods of refinement of retinal circuits. Further, loss of visually driven synaptic input prevents the structural maturation of AIS length shortening as well as increases synpo expression in RGCs.

## AIS of RGCs Have Similar Architectural Features as Principal Cortical Neurons

Here, we analyzed AIS positions along the axon by measuring the distance of the AIS to the RGC soma in adult animals.

We found that RGC AIS appear with a significant distance to the soma compared to pyramidal neurons in V1. AIS length and/or AIS position along the axon is proposed to impact neuronal excitability depending largely on the corresponding somatodendritic morphology of neurons (Gulledge and Bravo, 2016; Hamada et al., 2016; Kole and Brette, 2018). Thus, it is likely that the observed diversity in AIS position along the axon of RGCs in our study is linked to the variability of somatodendritic morphology within the different rodent RGC classes (Sun et al., 2002a,b; Sanes and Masland, 2015). As a result, optimal neuronal excitability could be adjusted by changing AIS length/position to individually limit or promote action potential generation.

By applying super resolution microscopy in neurons in vitro, the robust nanoscale organization along the AIS submembrane scaffold has been identified as a ∼190 nm periodic ring-like architecture formed by submembrane actin bands connected by longitudinal head-to-head βIV-spectrin subunits, which extend throughout the axon (Xu et al., 2013; D'Este et al., 2015; Leterrier et al., 2015). Despite the diverse AIS morphology in RGCs, our super resolution imaging data revealed a consistent nanoscale architecture along the AIS of RGCs. As seen in neurons in vitro, we observed a periodic arrangement of βIVspectrin with distances of ∼180 nm in RGC AIS. Within the AIS, we identified synpo-positive structures, corresponding to COs, localized underneath the membrane scaffold. Recently, it has been speculated that synpo-positive clusters within the AIS of cortical neurons in vitro and in vivo are localized at gaps in the AIS scaffold, which are deficient of ankyrin-G expression (King et al., 2014). These ankG and βIV-spectrin deficient sites are apparent in images acquired by super resolution imaging (STORM and STED) in AIS of neurons in vitro (D'Este et al., 2015; Leterrier et al., 2015). By applying SIM and SMLM, we observed such gaps in the RGC AIS scaffold, which were deficient of βIV-spectrin or ankG expression. However, these gaps were only occasionally filled with synpo-positive clusters and these clusters were often detected at sites in the AIS where no obvious gaps were visible. It is important to keep in mind that the super resolution methods applied here and in other studies have limitations. For example when using SMLM, the lack of threedimensional information might impact the interpretation of data. Moreover, a high density fluorescence labeling is required for SMLM to allow detection of positions of individual molecules for reconstructing a high resolution image (Klein et al., 2014). This could also impact the accuracy of measurements of positions of single molecules as we observed for measurements of distances between single ankG/βIV-spectrin signals (∼180 nm by STED vs. ∼200 nm by SMLM). A combination of methods should help to address the still unresolved question about 'gaps' in the AIS scaffold and possible colocalization of synpo clusters in these domains. Intriguingly, King and others speculated that gaps in the ankG/βIV-spectrin-actin scaffold are required to accommodate high density protein complexes that might be crucial for the assembly and functioning of axo-axonic GABAergic synapses at the AIS (Kosaka, 1980; Benedeczky et al., 1994; King et al., 2014). So far, the presence of such GABAergic synapses at RGC AIS is supported by an early electron microscopy study of the macaque monkey retina (Koontz, 1993). However, work supporting these findings in rodent RGCs is currently lacking. Furthermore, occasionally we observed synpo clusters that appeared as if they were partially located outside of the AIS. This phenomenon could have a technical explanation. Considering that ankG immunoreaction does not highlight the outside of the axonal membrane, but rather the underlying scaffold of the axon and hence, spans less axonal diameter, the actual complete axonal diameter may be underestimated. In addition, it is intriguing to speculate that the observation is further indication of a synpo/CO-subdomain in the AIS. There is evidence from hippocampal pyramidal neuron AIS that the synpo-positive clusters may represent 'spine-like' protrusions in the vicinity of – or as a direct contact site for – GABAergic synapses (Somogyi et al., 1983).

## Length of RGC AIS Correlates With Retinal Maturation During Postnatal Development

The AIS of RGCs is located proximally in the unmyelinated part of the axon and therefore, is separated spatially from the distal myelinated part (Hildebrand and Waxman, 1983; Koontz, 1993). The molecular composition of RGC AIS is comparable to that of other neurons across different species, and has been demonstrated by several studies (Hildebrand and Waxman, 1983; Carras et al., 1992; Koontz, 1993; Boiko et al., 2003; Van Wart et al., 2007; Zhang et al., 2015). Yet, the developmental maturation of AIS length in retinal neurons remained unexplored.

AIS length and location has been linked to the excitability of neurons (Grubb and Burrone, 2010; Kuba et al., 2010; Hamada et al., 2016; Jamann et al., 2018). Data support the hypothesis that longer AIS facilitate action potential generation and thus contribute to increased neuronal excitability. Interestingly, neuronal activity and excitability are inversely related, i.e., higher activity results in AIS shortening and lower activity leads to AIS elongation, a mechanism reminiscent of homeostatic plasticity (reviewed in Adachi et al., 2015; Wefelmeyer et al., 2016).

During cortical development, AIS undergo structural length maturation from increased AIS length at embryonic and early postnatal ages to a shortening at later postnatal stages and in adulthood (Cruz et al., 2009; Galiano et al., 2012; Fish et al., 2013; Gutzmann et al., 2014; Jamann et al., 2018). Here, we addressed the question whether similar developmental profiles apply for RGCs. Indeed, at postnatal ages around eye-opening (P10/15), AIS were longest with heterogeneous length distributions indicative of 'juvenile' AIS in V1 (Gutzmann et al., 2014). A characteristic feature of the developing retina is spontaneous periodic activity (reviewed in Firth et al., 2005). Spontaneous retinal waves emerge around E16.5, undergo different stages, and last until eye-opening (reviewed in Huberman et al., 2008). Vision in mice begins around P11 through naturally closed eye lids (Krug et al., 2001). The simultaneous presence of both spontaneous retinal activity and visually driven activity might provoke the increase in AIS length of RGCs in young animals. In turn, increased excitability is proposed to be acquired for early development in sensory cortices (Oswald and Reyes,

2008; Frangeul et al., 2017). It is conceivable that increased AIS length during retinal development leads to increased RGC excitability, which contributes to the refinement of axonal projections from the retina to the visual centers of the brain. Indeed, glutamate release from bipolar cells during spontaneous retinal waves from P10–12 regulates circuit development in the retina, and patterns of RGC activity propagating forward shape the wiring of circuits in the lateral geniculate nucleus, superior colliculus and in V1 (reviewed in Kerschensteiner, 2016). Interestingly, RGC activity is fundamental not only for proper development. Abnormalities in RGC activity were observed during development of pathologies and increased activity is discussed to decelerate the degenerative progression of retinal disease (Risner et al., 2018).

 The complete replacement of spontaneous activity with visually driven activity possibly triggered the decrease in AIS length in RGCs at P21, accompanied by a parallel homogenous AIS length distribution, which is characteristic for mature AIS in adult animals (Gutzmann et al., 2014). With proceeding development, AIS of RGCs decreased further in length, underlined by progressively more homogeneous AIS length distributions. Our findings indicate that AIS begin to mature when spontaneous retinal activity ends, suggesting that visual experience is important for the maturation of RGC AIS.

## Synpo Expression in RGC AIS Changes During Retinal Development and Stabilizes AIS Length Maturation

In rodent V1, synpo expression in the AIS begins postnatally and is most prominent in structurally mature AIS of pyramidal neurons during early critical periods of cortical development (Schlüter et al., 2017). In the present study, the first synpopositive clusters appeared in RGC AIS in P10 mice. Of note, maturation changed the size of synpo-positive clusters in RGC AIS, but the number of clusters per AIS was unaltered. In comparison to our study in the developing V1 (Schlüter et al., 2017), synpo expression was more constant during retinal development. The subset of AIS, which express synpo, amounted to ∼24 – 28% between the different ages and taking all RGC types into account. It should be pointed out that synpo clusters may possibly be associated with specific RGC classes, however, due to the overall scarcity of RGC<sup>A</sup> cells in our study, we cannot provide more detailed data at this point. The question whether synpo/CO segregates into specific RGC classes will have to be addressed in future studies. Interestingly, synpo-positive clusters within AIS were significantly different in size between P21 and P28 as well as P21 and P35. What could trigger the size reduction of synpo clusters during late postnatal development? One of the most important messengers for the induction of neuronal signaling during retinal development is Ca2<sup>+</sup> (Firth et al., 2005; Kerschensteiner, 2016). The CO and synpo have been implied in regulating local AIS Ca2<sup>+</sup> trafficking (Benedeczky et al., 1994; Sanchez-Ponce et al., 2012b; King et al., 2014). Ca2<sup>+</sup> sensitive channels (Inositol 1,4,5-trisphosphate receptors, ryanodine receptors) and Ca2<sup>+</sup> pumps (i.e., sarco/endoplasmic reticulum Ca2+-ATPase) associated with the CO are discussed to boost local cytosolic Ca2<sup>+</sup> transients during action potential firing through Ca2+-induced Ca2<sup>+</sup> release from internal Ca2<sup>+</sup> stores (Berridge, 1998; Segal, 2018). This Ca2<sup>+</sup> signaling amplification in turn could impact axonal membrane potential properties and neuronal excitability (Berridge, 1998; Segal, 2018), and thus AIS length.

In the developing retina, RGCs spontaneously fire periodic bursts of action potentials with accompanying large increases of intracellular Ca2<sup>+</sup> levels (reviewed in Firth et al., 2005). In mice, stage III – glutamatergic activity emerges from P10-14 (reviewed in Huberman et al., 2008; Kerschensteiner, 2016), declines around the time of eye opening as light-evoked signals begin to drive retinal activity, and are finally absent around P21 (Demas et al., 2003). The initial expression of synpo in RGC AIS at P10 may be linked to the appearance of stage III retinal glutamatergic waves and the imminent onset of vision. Early visual, experience along with the replacement of glutamatergic waves with light-evoked, signals may trigger a later decrease of sizes of synpo-positive clusters along with AIS length maturation of RGCs. Further, the density of synapses in the outer and IPL peaks around P21 (Xu and Tian, 2004). From P22 until P27, an increase of spontaneous excitatory and inhibitory postsynaptic currents emerges, which enhances RGC synaptic input more than fourfold (Tian and Copenhagen, 2001). This increase of synaptic input may change the firing properties of RGCs, which in turn might lead to the observed remodeling of synpo and the CO in retinal AIS.

In parallel with the emergence of synpo clusters in RGC AIS at P10, we observed a homogenous AIS length distribution compared to the entire AIS population in the retina at this age reflecting mature AIS (Gutzmann et al., 2014; Schlüter et al., 2017). The maturation of synpo-positive AIS early in development suggests that synpo presence in the AIS is a sign of structural maturity. Similar findings have been described for synpo expression in the AIS of pyramidal neurons during V1 development (Schlüter et al., 2017) or in dendritic spines (Mundel et al., 1997; Czarnecki et al., 2005).

#### Dark-Rearing Prevents Structural AIS Maturation and Remodels COs Within RGC AIS

Decreased neuronal activity caused by sensory deprivation during early postnatal development results in significant elongation of AIS, displaying characteristics observed in immature AIS. These findings suggest that increased sensory input and network activity leads to a structural maturation highlighted by AIS shortening during critical periods of development (reviewed in Jamann et al., 2018). To test whether this maturation is also a common developmental mechanism in RGCs, we investigated the structural length maturation of RGC AIS in dark-reared animals. Of note, AIS of dark-reared mice were ∼150% longer than AIS in control animals. The

average length as well as cumulative length distribution of AIS in dark-reared animals corresponded to that of juvenile AIS in young controls. These findings suggest that lack of visual input and decreased network activity in the retina leads to the prevention of structural AIS maturation in RGCs after eye-opening. Elongated AIS could maintain neuronal excitability at basal levels when sensory input is absent, which might reflect a homeostatic reaction for AIS under deprivation conditions.

Of note, the subset of RGCs that express synpo in their AIS, whose percentage was small throughout retinal development, were ∼120% higher in dark-reared animals. In all deprivation conditions, synpo-positive AIS were ∼110% longer than AIS in control animals of the same age and reached AIS length comparable to those observed in younger control animals. Further, sizes of synpo-positive clusters within RGC AIS were increased after dark-rearing. This could indicate that existing synpo-positive clusters fuse within the AIS after loss of visually driven synaptic input. The expansion of the ER most likely regulates the capacity of internal Ca2<sup>+</sup> stores (Sammels et al., 2010). Likewise, the formation of increased synpo-positive clusters is proposed to enhance Ca2<sup>+</sup> storage capacity in the spine apparatus in dendrites (Vlachos et al., 2013; Verbich et al., 2016) as well as in the CO in AIS (Schlüter et al., 2017) as a homeostatic response to loss of synaptic input. We speculate that both elongated AIS and expanded synpo-positive clusters within the AIS increase the excitability of RGCs as well as Ca2<sup>+</sup> currents in their AIS to compensate for the visual deprivation-triggered reduction in retinal network activity.

Finally, considering the significant heterogeneity of RGC classes, we would like to point out that further classification of RGC<sup>A</sup> into ON and OFF cells might be of interest since dark rearing might have differential effects on ON and OFF cells; it should silence ON cells and activate OFF cells, with presumably opposite AIS plasticity. Future studies in animal models more suited to discern a larger number of RGC classes will be able to shed light on this interesting question.

#### CONCLUSION

Our findings suggest that changes in synpo expression are linked to different stages of activity-driven processes in the developing retina. Moreover, we identified a possible role for both AIS and synpo/CO plasticity during homeostatic responses of visual input-deprived RGCs to reduced retinal network activity. Future studies could therefore focus on the subtypes of synpo-expressing RGCs, their intrinsic AIS plasticity and how it pertains to the development and maturation of functional visual circuits.

#### DATA AVAILABILITY

The datasets generated for this study are available on request to the corresponding author.

## ETHICS STATEMENT

All animal protocols were approved by the Medical Faculty Mannheim Animal Research Board, Heidelberg University, as well as the State of Baden-Württemberg, Germany, and were conducted in accordance with Heidelberg University Guidelines on the Care of Laboratory Animals.

#### AUTHOR CONTRIBUTIONS

AS, SR, and ME conceptualized and designed the study. AS, SR, DD, JMJ, SV, and JH contributed to data acquisition and programming. AS, SR, DD, JMJ, and ME analyzed the data. AS, SR, CS, and ME interpreted the data. AS, DM, and ME prepared the manuscript. All authors approved the final version to be published.

#### FUNDING

This work was supported by the ProRetina Foundation (Pro-Re/KP/Engelhardt.1-2014/2016 to ME) and the Deutsche Forschungsgemeinschaft (SFB1134/A03 to CS and ME; SFB1158/A08 and FOR2325 to DM). DM is a member of CellNetworks. DD was funded by the MD Fellowship program of the Medical Faculty Mannheim, Heidelberg University. This work was supported by the Core Facility Life Cell Imaging Mannheim (LIMA) at the CBTM (DFG-INST 91027/10-1 FUGG). We acknowledge financial support by the Deutsche Forschungsgemeinschaft within the funding program Open Access Publishing, by the Baden-Württemberg Ministry of Science, Research and the Arts and by the Ruprecht-Karls-University Heidelberg.

#### ACKNOWLEDGMENTS

We thank Prof. C. Cremer at the Kirchhoff Institute, Heidelberg University for usage of the SIM/SMLM setup, Dr. Sidney Cambridge, Department of Anatomy, Heidelberg University, for breeding pairs of the Thy1-EGFP mouse line, and Elisabeth Mertens for her technical support. We are indebted to Dr. Elisa D'Este, Max Planck Institute of Medical Research, Heidelberg for sharing her expertise in STED microscopy and to Johannes Roos, Institute of Neuroanatomy, Medical Faculty Mannheim, for providing the AISuite software for morphometrical analysis.

#### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fncel. 2019.00318/full#supplementary-material

#### REFERENCES

fncel-13-00318 July 27, 2019 Time: 14:57 # 17

**Conflict of Interest Statement:** SR joined DELMIC B.V. in the Netherlands, a company producing solutions for correlative light and electron microscopy, after completion of all work relating to the current study. Therefore SR declares no conflict of interest. JH works for Abberior Instruments GmbH, a company producing STED microscopes. DM is cofounder and shareholder of FundaMental Pharma GmbH, which has no conflict of interest with the present work.

The remaining authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2019 Schlüter, Rossberger, Dannehl, Janssen, Vorwald, Hanne, Schultz, Mauceri and Engelhardt. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

fncel-13-00407 September 4, 2019 Time: 17:3 # 1

# Excitability Tuning of Axons by Afterdepolarization

#### Haruyuki Kamiya\*

Department of Neurobiology, Hokkaido University Graduate School of Medicine, Sapporo, Japan

The axon provides a sole output of the neuron which propagates action potentials reliably to the axon terminal and transmits neuronal information to the postsynaptic neuron across the synapse. A classical view of neuronal signaling is based on these two processes, namely binary (all or none) signaling along the axon and graded (tunable) signaling at the synapse. Recent studies, however, have revealed that the excitability of the axon is subject to dynamic tuning for a short period after axonal action potentials. This was first described as post-spike hyperexcitability, as measured by the changes in stimulus threshold for a short period after an action potential. Later on, direct recordings from central nervous system (CNS) axons or axon terminals using subcellular patchclamp recording showed that axonal spikes are often followed by afterdepolarization (ADP) lasting for several tens of milliseconds and has been suggested to mediate postspike hyperexcitability. In this review article, I focused on the mechanisms as well as the functional significance of ADP in fine-scale modulation of axonal spike signaling in the CNS, with special reference to hippocampal mossy fibers, one of the best-studied CNS axons. As a common basic mechanism underlying axonal ADP, passive propagation by the capacitive discharge of the axonal membrane as well as voltage-dependent K <sup>+</sup> conductance underlies the generation of ADP. Small but prolonged axonal ADP lasting for several tens of milliseconds may influence the subsequent action potential and transmitter release from the axon terminals. Both duration and amplitude of axonal spike are subject to such modulation by preceding action potential-ADP sequence, deviating from the conventional assumption of digital nature of axonal spike signaling. Impact on the transmitter release is also discussed in the context of axonal spike plasticity. Axonal spike is subject to dynamic control on a fine-scale and thereby contributes to the short-term plasticity at the synapse.

Keywords: axon, action potential, afterdepolarization, propagation, short-term plasticity

## DYNAMIC TUNING OF AXON EXCITABILITY BY ADP

The axon carries neuronal information as a form of action potentials which reliably propagate for a long distance without attenuation (Debanne et al., 2011). A regenerative nature of spike generation provides digital property beneficial to reliable and ultrafast axonal signaling in the nervous system (Bean, 2007). Recent studies however, updated the classical view of digital axonal signaling to

Edited by:

Josef Bischofberger, University of Basel, Switzerland

#### Reviewed by:

Stefan Hallermann, Leipzig University, Germany Henrique Prado von Gersdorff, Oregon Health & Science University, United States Dmitri A. Rusakov, University College London, United Kingdom

\*Correspondence:

Haruyuki Kamiya<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Cellular Neurophysiology, a section of the journal Frontiers in Cellular Neuroscience

Received: 27 May 2019 Accepted: 26 August 2019 Published: 06 September 2019

#### Citation:

Kamiya H (2019) Excitability Tuning of Axons by Afterdepolarization. Front. Cell. Neurosci. 13:407. doi: 10.3389/fncel.2019.00407

**135**

fncel-13-00407 September 4, 2019 Time: 17:3 # 2

impart analog modification by the preceding neuronal activity (Zbili and Debanne, 2019). Such a use-dependent analog modification of axonal spike is possibly due to dynamic control of excitability of axon by the preceding neuronal activity for a short period up to tens to hundreds of milliseconds after generation of action potential (Gardner-Medwin, 1972; Zucker, 1974; Bucher and Goaillard, 2011). This post-stimulus change in the excitability of axon was mediated by ADP following action potential (**Figure 1A**), which may be important for temporal integration of axonal excitability and short-term plasticity of presynaptic transmitter release (Barrett and Barrett, 1982; Ohura and Kamiya, 2016). In this review, I focused on the recent progress in understanding the mechanisms as well as the functional significance of axonal afterdepolarization (ADP) in paired-pulse modulation and short-term synaptic plasticity.

#### COMPONENTS OF AXONAL ADP

ADP following action potentials is common process observed in the nervous systems in both vertebrate and invertebrate axons (Barrett and Barrett, 1982; Borst et al., 1995; Geiger and Jonas, 2000). A small but prolonged depolarization during ADP may important factor for activity tuning of axonal excitability as well as presynaptic transmitter release. **Figure 1A** shows the typical time course of ADP in hippocampal mossy fiber model (Engel and Jonas, 2005) calculated at negative membrane potentials (−100 mV) which shows similar time course with those recorded from mossy fiber boutons experimentally (Geiger and Jonas, 2000; Ohura and Kamiya, 2018b). This model assumes a Hodgkin Huxley-type gating adapted to channels those recorded in mossy fiber terminals and implemented with K<sup>+</sup> channel inactivation. There is a considerable number of studies addressing the mechanisms underlying ADP, and it seems to be reasonable to comprehend that axonal ADP consists of common basic mechanisms and of additional specific mechanisms to particular axons.

As a common mechanism underlying axonal ADP, passive propagation of upstream action potential via axon cable has been shown to consist of a basic component of ADP (Barrett and Barrett, 1982; Borst et al., 1995; David et al., 1995). Additional contribution of several specific mechanisms, e.g., activation of slow sodium current like resurgent or persistent sodium current (Kim et al., 2010; Ohura and Kamiya, 2018b), accumulation of potassium ions surrounding axons (Malenka et al., 1981; Kocsis et al., 1983; Meeks and Mennerick, 2004), or activation of autoreceptors of glutamate and GABA (Kamiya et al., 2002; Ruiz et al., 2003; Stell et al., 2007; Zorrilla de San Martin et al., 2017), have been also suggested.

#### PASSIVE PROPAGATION VIA AXON CABLE

Cable property of axon confers a delayed depolarization due to capacitive discharge from the upstream action potential. Such a passive propagating component consists of axonal ADP at

FIGURE 1 | Components of axonal ADP. (A) Basic components constituting axonal action potential-ADP sequence. Action potentials recorded from axon or the terminal are typically followed by prolonged ADP lasting for several tens of milliseconds. The left trace represents the time course of ADP in the hippocampal mossy fiber model calculated at negative membrane potentials of –100 mV. The right traces shows ionic components due to activation of voltage-dependent Na<sup>+</sup> (VNa, blue) and K<sup>+</sup> conductance (VK, green), the electrical component due to capacitive discharge (VCap, red) also substantially contribute to the prolonged ADP. VNa, VK, and VCap were calculated by subtraction of membrane potentials calculated by removal of Na<sup>+</sup> and/or K<sup>+</sup> conductance from the terminals. Sum of VNa, VK, and VCap, therefore, was identical with action potential-ADP sequence shown in the left trace. The inset represents the time-expanded traces showing timing and sequence of the onset of VNa, VK, and VCap. The electrical component (VCap) precedes the ionic components (VNa and VK) to trigger action potential at the downstream axons. (B) Passive propagation of upstream action potential. Using a model simulation of en passant axon (10 boutons spaced every 100 µm) mimicking the structure of hippocampal mossy fibers, the relative contribution of passive propagation to the downstream ADP was evaluated by removing voltage-dependent ionic conductance (gNa and gK) from 8th–10th boutons (Continued)

#### FIGURE 1 | Continued

fncel-13-00407 September 4, 2019 Time: 17:3 # 3

and axon, as shown in red. Brief current injection into the soma elicited action potential which propagates faithfully to the 7th bouton without attenuation. The amplitude of depolarization decreased successively from the 8th bouton, and the time course was slowed down along the distance due to filtering by axon cable. The decay of the peak amplitude along the distance was fitted by a single exponential curve with 53 µm for the distance with a reduction to 1/e (37%). (C) Boosting axonal ADP by slow Na<sup>+</sup> channels. In direct whole-cell recording experiment from hippocampal mossy fiber terminals, ADP is partly mediated by tetrodotoxin (TTX)-sensitive slow activating Na<sup>+</sup> channels. In the presence of TTX, action potentials and ADP were abolished, but brief current pulse injection restored sharp depolarization followed by slow relaxation (blue). Veratridine, an inhibitor of inactivation of Na<sup>+</sup> channels, enhanced the ADP and sometimes overlaid by multiple spiking elicited by a single stimulus in mossy fiber axons (red). Modified with permission from Ohura and Kamiya (2018b) and Kamiya (2019).

least in part. Consistent with this notion, ADP recorded from a calyx of Held axon terminals was shown to be unaffected by tetrodotoxin focally applied to the axon terminals (Borst et al., 1995), suggesting the possible axonal origin of ADP. Passive nature of ADP has also supported the finding that ADP and passive electrotonic response to step hyperpolarizing current injection showed similar time courses in motor axons (Barrett and Barrett, 1982) or in hippocampal mossy fiber axons (Ohura and Kamiya, 2018b).

To evaluate quantitative contribution of passive propagation in action potential-ADP sequence, time courses of the ionic components due to activation of voltage-dependent Na<sup>+</sup> and K<sup>+</sup> channels (VNa and VK) as well as electrical components due to capacitive discharge (VCap) are compared (**Figure 1A**) in the hippocampal mossy fiber model used in our recent simulation study (Kamiya, 2019). The amplitude of VCap is more than onethird of VNa, suggesting a substantial contribution of passive electrical components in ADP, in agreement with the experiment of brief current injection in the presence of tetrodotoxin, a blocker of voltage-dependent Na<sup>+</sup> channels (Ohura and Kamiya, 2018b). The onset of VCap preceded VNa to trigger action potentials as shown in the inset with an expanded time scale.

It should be noted that passive propagation may distribute over a relatively long distance. The length constant of the axon was estimated as 455 µm in layer V pyramidal neurons in ferret cortex (Shu et al., 2006), 450 µm in rat hippocampal granule cell (Alle and Geiger, 2006), and 121 µm in cultured rat Purkinje cell (Zorrilla de San Martin et al., 2017). Although the length constant evaluated by long current pulse injection was estimated as 171 µm in our model of hippocampal mossy fibers (Kamiya, 2019), action potentials decline with shorter space constant of 53 µm for decay to 1/e (**Figure 1B**) possibly reflecting steeper filtering of fast voltage transient during action potentials. Passive propagation filtered by axon cable may thereby substantially impact the time course of action potential-ADP sequence recorded from the downstream axon.

It is worth considering whether subthreshold fluctuation of somatic membrane potentials occurring in vivo may also passively propagate to the axon. Long-range propagation of somatic depolarization into the axon has been demonstrated for hippocampal mossy fibers (Alle and Geiger, 2006) and for cortical pyramidal cell axons (Shu et al., 2006). In both studies, axonal depolarization enhanced transmitter release from the axon terminals, while the effect was limited to the proximal portion of the axons (see also Scott et al., 2008) due to passive attenuation of depolarization with the distance from the soma. On the contrary, ADP following action potentials is expected to equally distribute along the course of the axon, since action potentials propagate without attenuation. Therefore passive propagating components of axonal ADP is not depending on the distance from the soma and differed from the subthreshold somatic depolarization in this point.

Contribution of passive propagation is important for transmitter release not only from en passant boutons like the hippocampal mossy fibers, but also from bouton terminaux like calyx of Held, since it has been shown that Na<sup>+</sup> channels are excluded from the terminals but are expressed only in the axonal heminode in the distal axon (Leão et al., 2005). The depolarization at the active zone, which is directly related to Ca2<sup>+</sup> entry responsible for transmitter release, is therefore mostly reflecting passive propagation from the heminode region to the terminal boutons.

## VOLTAGE-DEPENDENCY OF AXONAL ADP

Quite puzzling observations are ADP recorded from axons or the terminals showed clear voltage-dependency. The size of ADP decreased upon depolarization of the initial membrane potentials (Begum et al., 2016; Sierksma and Borst, 2017; Ohura and Kamiya, 2018b), and sometimes reversed in polarity at more positive membrane potentials. It was also demonstrated that hyperpolarizing current injection decreased ADP (Kim et al., 2010). These findings are difficult to interpret with the passive nature of capacitive discharge of the axonal membrane, which is fundamentally voltage-independent. Therefore, the additional contribution of voltage-gated conductance which provides voltage-dependency to axonal ADP (Kamiya, 2019) must be taken into consideration.

#### K <sup>+</sup> CHANNELS SHAPE THE INITIAL PHASE OF AXONAL ADP

As additional conductance that potentially confers voltagedependency to axonal ADP, the contribution of voltagedependent K<sup>+</sup> conductance, which predominantly mediates fast repolarization of action potential (Storm, 1987; Dodson et al., 2003; Wissmann et al., 2003), was suggested. The time course of K<sup>+</sup> conductance somewhat outlasts the duration of action potentials and necessarily contributes to the subsequent ADP time course. Consistent with this notion, voltage-dependent K<sup>+</sup> conductance shapes a characteristic breakpoint at the initial phase of axonal ADP, possibly due to superimposed hyperpolarizing K <sup>+</sup> conductance on the passive depolarizing component. Sum of voltage-independent passive component and voltagedependent K<sup>+</sup> channel component with the various combination fncel-13-00407 September 4, 2019 Time: 17:3 # 4

in hippocampal mossy fiber model nicely reconstruct the characteristic time course as well as typical voltage-dependency of action potential-ADP sequence (Kamiya, 2019). It seems to be reasonable to speculate that various reported values of apparent reversal potential of axonal ADP (Begum et al., 2016; Sierksma and Borst, 2017; Ohura and Kamiya, 2018b) reflect the various contribution of passive propagation and voltage-dependent K+ channel components.

## SLOW Na<sup>+</sup> CURRENT BOOSTS ADP IN SOME AXONS

In addition to the common basic mechanisms of passive propagation and voltage-dependent K<sup>+</sup> conductance, the contribution of slow Na<sup>+</sup> current at a certain type of axon has been suggested. ADP recorded from soma sometimes are mediated by slow voltage-dependent Na<sup>+</sup> currents such as persistent-type INaP or resurgent-type INaR (Raman and Bean, 1997; Yue and Yaari, 2004; Yue et al., 2005; D'Ascenzo et al., 2009). Using direct recording from the calyx of Held axon terminals, thereafter, it has been shown that the resurgent-type Na<sup>+</sup> current INaR shapes slow time course of axonal ADP (Kim et al., 2010). In contrast, it was reported that ADP was unaffected by local application of tetrodotoxin surrounding the recorded terminals at the same axon terminals (Borst et al., 1995), although the reason of the different results in these studies is unclear. This discrepancy may be explained by the finding that Na<sup>+</sup> channels are not located in the calyx terminal but in the distal heminode region of the axon (Leão et al., 2005). Recently we also have reported that ADP at hippocampal mossy fiber terminals is also partially mediated by voltage-dependent Na<sup>+</sup> current (Ohura and Kamiya, 2018b) since it was suppressed by a Na<sup>+</sup> channel blocker tetrodotoxin and was enhanced by an inhibitor of inactivation of Na<sup>+</sup> channel veratridine (**Figure 1C**), as shown for ADP at the calyx of Held terminals (Kim et al., 2010). These slow Na<sup>+</sup> channels raise axonal excitability for a while and help to support faithful spiking of axons during repetitive stimuli. Their molecular identity, as well as the precise subcellular localization of the axonal slow Na<sup>+</sup> channels, remain to be determined. It is also uncertain whether this mechanism is generally applicable to other types of axons in different brain regions.

## ACCUMULATION OF K<sup>+</sup> SURROUNDING AXONS

It was reported that stimulation of parallel fiber axons in rat cerebellum cause elevation of extracellular K<sup>+</sup> concentration surrounding axons and caused prolonged depolarization and hyperexcitable period lasting for a hundred of milliseconds (Malenka et al., 1981; Kocsis et al., 1983). It is intriguing to speculate that action potential in axon or EPSPs in postsynaptic neurons caused elevation of extracellular K<sup>+</sup> concentration in the extracellular space and axon to depolarize by elevated K<sup>+</sup> concentration surrounding axons. It will be worth testing the possible contribution of astroglia in shaping axonal ADP since local extracellular K<sup>+</sup> buffering depends mainly on astroglia. It is also intriguing to test the roles of gliotransmitters in shaping axonal ADP, since glutamate released from astroglia was shown to broaden action potentials locally (Sasaki et al., 2011). However, the increase in elevated levels of K<sup>+</sup> was observed only when strong and repetitive stimuli were given. The single shock-induced K<sup>+</sup> increase was detectable only when a K<sup>+</sup> channel blocker 4-AP was applied. Therefore, it seems to be that this mechanism may not contribute to the generation of ADP physiologically by single action potential at single axon.

## CONTRIBUTION OF AUTORECEPTOR ACTIVATION IN AXONAL ADP

In some specific axons, it has been shown that activation of autoreceptors of GABA (Pouzat and Marty, 1999; Zorrilla de San Martin et al., 2017) participates in axonal ADP. Activation of axonal GABAA-autoreceptors at cerebellar interneuron axons causes excitatory GABAergic autoreceptor currents (Pouzat and Marty, 1999), possibly due to higher chloride concentration in axoplasm, and facilitate transmitter release and increase neuronal firing rate (Mejia-Gervacio and Marty, 2006). Similar depolarizing autoreceptor current was also reported for cultured Purkinje cell axons (Zorrilla de San Martin et al., 2017). It was also suggested that kainate-type glutamate receptors may assist ADP in hippocampal mossy fiber axons recorded optically using voltage-sensitive dye (Kamiya et al., 2002). It should be noted that autoreceptor activation surely contributes to a certain subset of axons in the nervous system, this is not a common mechanism for all types of the axon, but a relatively rare and specific contribution to axonal ADP.

### IMPACT OF AXONAL ADP ON SUBSEQUENT ACTION POTENTIAL AND TRANSMITTER RELEASE

Slow time course of axonal ADP implies that the excitability of the axon is modulated cumulatively during repetitive stimuli at short intervals. The small and slow depolarization of ADP may affect the states of the voltage-dependent Na<sup>+</sup> and K<sup>+</sup> channels and thereby potentially modify the subsequent action potentials. Using whole-bouton recording from mossy fiber terminals, we recently have reported (Ohura and Kamiya, 2018b) that the peak height of the action potentials was almost unchanged, although the amplitudes of the action potentials measured from the elevated initial membrane potential were reduced in paired-pulse stimuli at short intervals (**Figure 2A**). Elevated membrane potentials by ADP may facilitate steadystate inactivation of voltage-dependent Na<sup>+</sup> channels (Engel and Jonas, 2005; Rama et al., 2015), and thereby may slow the rising phase of action potentials. Consistent with this, axonal spike recorded extracellularly from single mossy fiber terminal, which is expected to reflect the first derivative of action potential recorded intracellularly (Meeks et al., 2005), displayed short-term depression of the amplitude at short fncel-13-00407 September 4, 2019 Time: 17:3 # 5

intervals (Ohura and Kamiya, 2018a) as shown in **Figure 2B**. Inactivation of Na<sup>+</sup> channels was also suggested for mediating adaptive broadening of spike initiation site during somatic depolarization, as demonstrated by simultaneous soma and axon recordings from hippocampal mossy fibers (Scott et al., 2014). It was also demonstrated that the preceding ADP also enhanced inactivation of K<sup>+</sup> channels, leading to a usedependent broadening of action potentials (Geiger and Jonas, 2000; Kole et al., 2007).

Voltage-dependent Ca2<sup>+</sup> channels are also affected by the preceding ADP. Modulation of presynaptic Ca2<sup>+</sup> current would be expected to modulate transmitter release and short-term synaptic plasticity (Bischofberger et al., 2002; Li et al., 2007; Zorrilla de San Martin et al., 2017). Although the detailed biophysical mechanism remains to be clarified, facilitation of Ca2<sup>+</sup> current consequently enhances the subsequent transmitter release from the axon terminals at the culture Purkinje cell axon terminals (Zorrilla de San Martin et al., 2017) and hippocampal mossy fiber terminals (Ohura and Kamiya, 2018b). Impacts on the downstream synaptic transmission is a small but non-negligible role of axonal ADP since the amount of transmitter release is steeply dependent on the amount of Ca2<sup>+</sup> entry during an action potential supralinearly (Zucker and Regehr, 2002). In fact, it has been shown that subthreshold depolarization of the calyx of Held raised the Ca2<sup>+</sup> levels by weak activation of P/Q-type Ca2<sup>+</sup> channels and enhanced transmitter release from the terminals (Awatramani et al., 2005).

by the paired stimuli at short intervals. (B) Paired-pulse depression of axonal spikes recorded from single mossy fiber boutons by loose-patch clamp recordings. The

amplitude of the second spike was slightly reduced than the first spike at short intervals. Modified with permission from Ohura and Kamiya (2018a,b).

## CONCLUSION

fncel-13-00407 September 4, 2019 Time: 17:3 # 6

Recent studies using the direct electrophysiological recordings from axons or axon terminals in the central nervous system have revealed that the excitability of axons is regulated more dynamically than previously thought. In this review article, advances in the understanding of the mechanisms and the functional significance of ADP following axonal action potentials were summarized. It has been demonstrated that electrical signals passively propagate for hundreds of micrometers, and consequently, the passive propagating component due to the capacitive discharge of axons substantially contributes to the propagating action potential-ADP sequence. The changes in the excitability by axonal ADP last for tens or hundreds of milliseconds and therefore may play an important role in temporal integration of neuronal activity and short-term synaptic

### REFERENCES

plasticity. Such a fine-scale dynamics of axonal excitability play pivotal roles in fine-tuning and temporal integration of neuronal network functions.

#### AUTHOR CONTRIBUTIONS

HK conceptualized and designed the study and drafted the manuscript.

## FUNDING

This study was supported by Grant-in-Aid for Scientific Research (KAKENHI) from the Japan Society for the Promotion of Science (18K06514 to HK).

fncel-13-00407 September 4, 2019 Time: 17:3 # 7

**Conflict of Interest Statement:** The author declares that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2019 Kamiya. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Large-Scale Mapping of Axonal Arbors Using High-Density Microelectrode Arrays

Torsten Bullmann1,2,3, Milos Radivojevic<sup>4</sup> , Stefan T. Huber <sup>1</sup> , Kosmas Deligkaris 1,5 , Andreas Hierlemann<sup>4</sup> \* and Urs Frey 1,4,6

 *RIKEN Quantitative Biology Center, RIKEN, Kobe, Japan, <sup>2</sup> Graduate School of Informatics, Kyoto University, Kyoto, Japan, Carl Ludwig Institute for Physiology, University of Leipzig, Leipzig, Germany, <sup>4</sup> Department of Biosystems Science and Engineering, ETH Zurich, Basel, Switzerland, <sup>5</sup> Graduate School of Frontier Biosciences, Osaka University, Osaka, Japan, MaxWell Biosystems AG, Basel, Switzerland*

Understanding the role of axons in neuronal information processing is a fundamental task in neuroscience. Over the last years, sophisticated patch-clamp investigations have provided unexpected and exciting data on axonal phenomena and functioning, but there is still a need for methods to investigate full axonal arbors at sufficient throughput. Here, we present a new method for the simultaneous mapping of the axonal arbors of a large number of individual neurons, which relies on their extracellular signals that have been recorded with high-density microelectrode arrays (HD-MEAs). The segmentation of axons was performed based on the local correlation of extracellular signals. Comparison of the results with both, ground truth and receiver operator characteristics, shows that the new segmentation method outperforms previously used methods. Using a standard HD-MEA, we mapped the axonal arbors of 68 neurons in <6 h. The fully automated method can be extended to new generations of HD-MEAs with larger data output and is estimated to provide data of axonal arbors of thousands of neurons within recording sessions of a few hours.

Keywords: axons, high-density microelectrode array, extracellular electrical field, action potential, axonal arborizations, action potential propagation, high-throughput screening

#### INTRODUCTION

The classical view of axons is that of mere transmission cables (Hodgkin and Huxley, 1952), while dendrites integrate distinct synaptic inputs (Spruston, 2008), and learning and memory are perceived to be a consequence of synaptic plasticity (Redondo and Morris, 2011). Recent data, however, indicate that the functional capacity of axons may be much more complex (Debanne, 2004; Ohura and Kamiya, 2016; Rama et al., 2018).

According to the classical theory describing giant squid axons (Hodgkin and Huxley, 1952), voltage-gated sodium and potassium channels support self-sustained action potentials that propagate along the axon. That is also true for mammalian axons, but at least two additional types of cationic channels have been described. These channels are activated by G-protein dependent receptors or hyperpolarization and modify the shape and propagation of the action potentials (Elgueta et al., 2015; Ko et al., 2016).

Axons can be surrounded by a myelin sheet that leads to saltatory conduction of action potentials (Tasaki, 1939), instead of the continuous conduction that has been observed in nonmyelinated axons, such as the giant squid axons (Hodgkin and Huxley, 1952). Mammalian axons

#### Edited by:

*Dominique Debanne, INSERM U1072 Neurobiologie des canaux Ioniques et de la Synapse, France*

#### Reviewed by:

*Adelaide Fernandes, University of Lisbon, Portugal Valentina Carabelli, University of Turin, Italy*

#### \*Correspondence:

*Andreas Hierlemann<EMAIL_ADDRESS>
#### Specialty section:

*This article was submitted to Cellular Neurophysiology, a section of the journal Frontiers in Cellular Neuroscience*

Received: *23 May 2019* Accepted: *20 August 2019* Published: *06 September 2019*

#### Citation:

*Bullmann T, Radivojevic M, Huber ST, Deligkaris K, Hierlemann A and Frey U (2019) Large-Scale Mapping of Axonal Arbors Using High-Density Microelectrode Arrays. Front. Cell. Neurosci. 13:404. doi: 10.3389/fncel.2019.00404* also show considerable variations in their diameter (Gaussian distribution around a peak at 200 nm diameter) and their number of branch points and varicosities, which affect the conduction velocity of action potentials. Differences in axon lengths have been found to produce defined temporal delays for spike trains for coincidence detection in the auditory system (Seidl et al., 2010; Stange-Marten et al., 2017). Furthermore, axonal delays in the neocortex show sub-millisecond precision (Swadlow, 1994), which is compatible with mechanisms of spike-timing-dependent synaptic plasticity (Dan and Poo, 2004). These characteristics lead, according to computational studies (Izhikevich, 2006), to the emergence of precisely timed firing patterns.

Early computational studies have suggested that some axonal arbors can act as filters for spike patterns by the selective failure of action potential propagation (Lüscher and Shiner, 1990). Whereas cerebellar mossy fibers in the cerebellum can reliably transmit spike trains up to 1.6 kHz (Ritzau-Jost et al., 2014), occasional failures have been observed in the axons of CA3 pyramidal neuron at firing frequencies of 30–40 Hz (Meeks and Mennerick, 2007). Failure depends on the diameter and the branching morphology as well as on the activation of presynaptic A-type potassium channels. The morphology and molecular composition of the individual axonal arbors has been found to influence the timing and shape of the action potential (Bischofberger et al., 2002; Alle and Geiger, 2006; Cho et al., 2017) arriving at the presynapse, which affects synaptic transmission and plasticity. For example, broadening of action potentials due to slow inactivation of voltage-gated potassium channels during high-frequency spike trains was found to facilitate synaptic release (Geiger and Jonas, 2000). Furthermore, axonal signaling and synaptic connectivity have been found to constitute important parameters in induced-pluripotent-stem-cell (iPSC) models of Parkinson's (Kouroupi et al., 2017) and amyotrophic lateral sclerosis (Wainger et al., 2014).

The understanding of information processing in axons is a fundamental question in neuroscience; however, the availability of experimental data is severely limited due to the small axon diameter. Most data sets have been gathered by performing patch clamp recordings of axonal membranes, at axon terminals and boutons, at the intact axon shaft or at the axonal bleb that is formed upon cutting axons (Ohura and Kamiya, 2016). However, these patch-based methods are not very well-suited to track the propagation of action potentials at more than two sites or even across the full axonal arbor. Moreover, the patch recording time is limited to a few hours while axonal recording is a serial process and has to be done axon after axon. Finally, patchbased methods constitute endpoint measurements and following the development of axonal arbors over extended periods is not possible. Another possibility to study axons includes the use of imaging tools and optogenetics (Chen et al., 2013), and it has been shown that imaging of single axon terminals is possible (Hoppa et al., 2014). However these methods are still limited in signal-to-noise ratio as well as in temporal resolution (Emmenegger et al., 2019).

A viable alternative to the methods described above includes the use of high-density microelectrode arrays (HD-MEAs), based on complementary-metal-oxide-semiconductor (CMOS) technology (Eversmann et al., 2003; Berdondini et al., 2009; Frey et al., 2010; Ballini et al., 2014; Bertotti et al., 2014; Viswam et al., 2016; Tsai et al., 2017). HD-MEAs can be used to capture neuronal activity at high temporal resolution across spatial scales, including networks, dendrites and most importantly, axons (Obien et al., 2015). CMOS-based HD-MEAs have been used to study the action potential propagation along axonal arbors (Bakkum et al., 2013) and the initiation of action potentials at the axon initial segment (Bakkum et al., 2019), as well as to track single action potentials along axons (Radivojevic et al., 2017) and stimulate single axon initial segments (Ronchi et al., 2019). Furthermore, HD-MEA recordings can be combined with classical patch clamp recording to study postsynaptic currents in response to pre-synaptic stimulation (Jäckel et al., 2017).

The objective of this work was to provide a method for highthroughput scanning of axonal arbors and mapping their axonal delays with minimal or no need to adjust parameters for the detection of axonal signals. Using a standard HD-MEA (Frey et al., 2010), we mapped the axonal arbors of more than 68 neurons in <6 h.

## MATERIALS AND METHODS

#### Animal Use

Timed pregnant rats (Wistar) were obtained from a commercial vendor (Nihon SLC, Japan). Animals were sacrificed on the day of arrival to obtain embryos for primary neuron cultures. All experimental procedures on animals were carried out in accordance with the European Council Directive of 22 of September 2010 (2010/63/EU) and have been approved by the local authorities in Japan (Animal Care and Use Committee of RIKEN; QAH24-01).

## High-Density Microelectrode Array (HD-MEA)

Sub-cellular resolution extracellular recordings were obtained using a CMOS-based HD-MEA (Frey et al., 2010) with 11,011 electrodes, arranged in a hexagonal pattern and featuring an electrode density of 3,150 electrodes/mm<sup>2</sup> . The culture chamber of the HD-MEA was prepared as described before (Heer et al., 2007) with minor modifications: after attaching the chamber ring (polycarbonate, 19 mm inner diameter, 8 mm high) using epoxy resin (EPO-TEK 301-2, Epoxy Technology Inc.), GlobTop (G8345D-37, Namics Inc.) was used to cover the bond wires while keeping the electrode area clean, and the remaining area was covered by a thin film of PDMS (Sylgard 184, Dow Corning). Platinum black was electrochemically deposited (Marrese, 1987) [with modifications of the original procedure (Heer et al., 2007)] on the electrodes to decrease their impedance in order to improve signal-to-noise characteristics (Viswam et al., 2014). Before plating, the surface of the HD-MEAs was rendered hydrophilic by oxygen-plasma treatment (40 s, 20 W), incubated for 4 h with of 50µg/ml Poly-D-Lysine (Sigma-Aldrich, P7280) in PBS, washed twice with aqua dest and air-dried for 1 h.

#### Primary Neuron Cultures

Adult rats were anesthetized with isofluorane and killed using a guillotine. The embryos were removed from the uterus and decapitated. Their neocortex was dissected in ice-cold dissection medium (HBSS without Ca2<sup>+</sup> and Mg2+; Gibco, NO.14175) and incubated for 20 min at 37◦C in Trypsin/EDTA (Sigma-Aldrich). After washing twice with plating medium (Neurobasal A, supplemented with 10% Fetal bovine serum, 2% B27 Supplement, 1:100 GlutaMax, all from Gibco, Japan, and 10µg/ml Gentamicin, Sigma-Aldrich, Japan), the tissue was mechanically dissociated, passed through a 40µm nylon mesh, and centrifuged 6 min at 200 g. The supernatant was removed; the cells were suspended and counted. A 20 µl drop containing 10,000 cells was placed on the electrode area of the HD-MEA in the middle of the culture chamber. The cultures were covered with a membrane, permeable to gas but not to water vapor, and placed in a standard incubator (37◦C, 5% CO2, 80% relative humidity). The neurons were allowed to settle and attach to the surface during 30 min. Thereafter, the culture chamber was filled with 600 µl serumfree, astrocyte-conditioned DMEM/Hams's F12 medium (Nerve Culture Medium, Sumitomo, Japan, #MB-X9501). Medium was completely exchanged with 600 µl conditioned medium after 4 days and then every 7 days until day 17 in vitro.

#### Recordings and Spike Event Detection

For recording, HD-MEAs were placed in a bench-top incubator (TOKAI HIT, Japan, INU-OTOR-RE) with temperature control, and 5% CO<sup>2</sup> was supplied by a gas-mixer and humidified by a water bath. In order to avoid the evaporation of medium during prolonged recording intervals, the water bath and the lid temperature set point was set 1K and 3K above the sample temperature set point, which was 35◦C. The HD-MEA recordings were performed using custom scripts, written in LabView (National Instruments, US), Matlab (Mathworks, US), C++ and Python running on a standard PC with a Linux operating system. Data underwent lossless data compression and were directly stored on a server on the local LAN.

Offline analysis of the recordings included filtering, event detection and averaging. First, a band-pass filter (2nd order Butterworth filter, 100–3,500 Hz) was used to remove slowly changing field potentials as well as high frequency noise. The remaining (background) noise was characterized by the median absolute deviation (MAD), which is resilient to outliers in the data but represents a consistent estimator of the standard deviation, <sup>s</sup><sup>V</sup> <sup>=</sup> 1.4826 MAD Vsig . Using a voltage-threshold method for event detection (Lewicki, 1998), negative signal peaks below a threshold of Vthr = 5s<sup>V</sup> (Vthr > 50 µV in all cases) were identified (**Figure 1C**). To avoid multiple detection of the same spike, successive events within <0.5 ms were discarded.

#### Mapping of Axonal Initial Segments

To initially identify the location of axonal initial segments (AISs), the whole array was scanned using configurations in which non-overlapping blocks of 6 × 17 electrodes (**Figure 1D**) were connected to the amplifiers through the switch matrix (Frey et al., 2010) (schematic in **Figure 1B**). After recording for 30 s from every electrode in the array, spike detection was performed for each electrode and taking the median value summarized the amplitude of the negative peaks. The median of the negative peak for each electrode was plotted as a map showing some areas with large negative peaks (**Figure 2A**). Such local minima were assumed to indicate the putative (proximal) AIS locations (Bakkum et al., 2019). We did not distinguish between inhibitory and excitatory neurons according to the AIS waveform (Mita et al., 2019).

#### Mapping of Electrical "Footprints"

To map the electrical "footprint" (spatial distribution of extracellularly measured electrical potentials obtained with the densely packed electrodes) of neuronal units and their axonal arbors over the entire array, we used a series of configurations in which so called "fixed electrodes" were always connected to the amplifiers through the switch matrix (Frey et al., 2010), whereas the remaining "variable electrodes" were connected in sequential configurations (schematic in **Figure 1E**). After recording the spontaneous activity for 115 s for each of these configurations, spike sorting was performed and the electrical "footprint" was calculated using the spike-triggered average as previously described Bakkum et al., 2013; Müller et al., 2015; Deligkaris et al., 2016; Radivojevic et al., 2016, 2017; Jäckel et al., 2017.

To perform spike-triggered averaging we need to reliably record spiking activity of the neurons. Therefore, we selected the fixed electrodes at the putative locations of the AISs but imposed a spatial restriction in order to not record the same neuron twice. For selection of fixed electrodes, all electrodes were ranked according to their median negative peak amplitude (see previous section). The electrode with the highest rank was selected, afterwards all electrodes in its proximity (within 100µm distance) were discarded from the list, and the procedure was repeated.

## Spike Sorting