There are three main different ways to achieve genetic targeting: (1) conventional/bacterial artificial chromosome (BAC) transgene (Schmidt et al., 2013), (2) gene knock-in (Taniguchi et al., 2011), and (3) gene/enhancer trapping (Leighton et al., 2001; Kelsch et al., 2012). Conventional or BAC transgene approaches use a transgene cassette, or BAC, to introduce the promoter/enhancer regions of the gene of interest to drive expression of a reporter which mimics endogenous expression patterns. They are randomly inserted in the genome, which may cause variations in expression patterns among founder lines. Gene knock-in relies on inserting the reporter gene in the endogenous location of the gene of interest to fully recapitulate endogenous expression patterns, however it is technically more challenging and expression of the target gene itself may be altered even using an internal ribosome entry site (IRES) sequence. In enhancer trapping, a reporter driven by a minimal promoter is randomly inserted in the genome, and local enhancers near the insertion site then determine expression of the reporter.

Discovery of green fluorescent protein (GFP) (Tsien, 1998) in 1962 was a significant milestone for cell and molecular biology, and it was granted the Nobel Prize in Chemistry in 2008 (Weiss, 2008). It was not long before GFP was used as a marker for gene expression in eukaryotes (Chalfie et al., 1994). Since then, there has been an explosion of its spectral variants with a wide variety of brightness and colors (Shaner et al., 2005). Transgenic mice expressing GFP under control of the cytomegalovirus (CMV) promoter allowed study of the central nervous system at a level of single individual neuron resolution (van den Pol and Ghosh, 1998). Spectral variants of GFP (yellow, YFP; cyan, CFP; and red, RFP; collectively called XFP) expressed in transgenic mice under the control of Thy-1 promoter allowed imaging and identification of numerous subsets of neurons (Feng et al., 2000). More recently, using a cre/lox recombination system to express multiple distinct colors in diverse genetically labeled neurons gave rise to the "Brainbow" mice (Livet et al., 2007; Cai et al., 2013).

Numerous reporter mouse lines have been generated using genetic targeting methods and extensively used to study various cell types in the mouse CNS, such as layer V pyramidal neurons (Feng et al., 2000; Yu et al., 2008), cortical interneurons (Chattopadhyaya et al., 2004; Lopez-Bendito et al., 2004; Taniguchi et al., 2011), basal ganglia (Gerfen et al., 2013), spinal cord motor neurons and interneurons (Wichterle et al., 2002; Miles et al., 2004; Chang and Martin, 2011). Of these, Thy-1 XFP mice are of particular interest as they allow analysis of labeled cells at a very fine detail, such as dendritic spine dynamics *in vivo* (Grutzendler et al., 2002; Oray et al., 2004; Dombeck et al., 2007). Moreover, they have been used to study mouse models of neurodegenerative diseases at a cellular level, such as dendritic spine loss in triple transgenic Alzheimer's disease mice (Bittner et al., 2010) and motor neuron pathology in the experimental autoimmune encephalomyelitis, and mouse model of multiple sclerosis (Bannerman et al., 2005). Thy1-YFP mice have extensively been used to study motor systems, not only SMN in mouse models of ALS (Schaefer et al., 2005; Wong et al., 2009), but also CSMN (Richter and Roskams, 2009) and their axons in spinal cord injury (Bareyre et al., 2005) and an ALS mouse model (Ozdinler et al., 2011).

Another reporter line extensively used to study motor systems is HB9-GFP, which labels about 90% of SMN in the developing embryo (Wichterle et al., 2002) but only about half of large Choline acetyl transferase (ChAT)+ SMN in the ventral horn of the adult (Chang and Martin, 2011). γ-SMN innervating muscle spindles are GFP− (Shneider et al., 2009). HB9-GFP mice have been used to follow differentiation of mouse embryonic stem cells into motor neurons (Wichterle et al., 2002; Wu et al., 2012). HP9-GFP+ embryonic stem cell-derived SMN co-cultured with astrocytes expressing hSOD1G93A displayed increased cell death and shorter axon length (Dodge et al., 2008). HB9-GFP BACs have also been used to characterize human embryonic stem cellderived motor neuron lineages (Placantonakis et al., 2009) or facilitate FACS purification of human spinal motor neurons from embryonic stem cells (Singh Roy et al., 2005). Human embryonic stem cell-derived motor neurons identified by HB9-GFP expression showed that SMN with hSOD1 mutations displayed reduced survival and shorter axons (Karumbayaram et al., 2009). Primary and embryonic stem cell-derived motor neurons from HB9-GFP mice have been used to study neurodegenerative properties of human SOD1 mutations, showing glia and astrocytes with SOD1 mutations induce neurodegeneration of co-cultured SMN even when they do not carry the mutation themselves (Di Giorgio et al., 2007; Nagai et al., 2007). HB9-GFP mice have been used to study (1) motor axon guidance *in vivo* by Semaphorin signaling (Huber et al., 2005), (2) *in vitro* by GDNF chemoattraction and ephrinA signaling (Dudanova et al., 2010) during normal development, (3) targeting of regenerating motor axons *in vivo* by polysialylated NCAM in the adult (Franz et al., 2005), and (4) motor axon development *in vivo* in mouse models of disease such as SMA (McGovern et al., 2008).

Gene Expression Nervous System Atlas (GENSAT) database has been invaluable in providing a detailed library of hundreds of distinct, genetically defined cell populations from engineered mice utilizing BAC (Gong et al., 2003, 2007; Doyle et al., 2008; Schmidt et al., 2013). To date, there are almost 400 publications that use GENSAT mice or BACs (www*.*GENSAT*.*org). Using this approach, we generated the UCHL1-eGFP mice in which the UCHL1 gene promoter was used to drive eGFP expression (**Figure 3**) (Yasvoina et al., 2013). CSMN identity of eGFP+ neurons was confirmed by retrograde labeling, molecular marker expression profile, electrophysiology, cortical circuit mapping, and mouse genetics studies. CSMN in the motor cortex and their projections were genetically and stably labeled by GFP expression from P0 to P800. In the spinal cord, almost all ChAT+ SMN were eGFP+ at birth but by P30 eGFP expression became restricted to a mixture of small α- and γ-SMN that were resistant to degeneration in motor neuron diseases, such as ALS. Crossing this reporter mouse with hSOD1G93A ALS mouse model (Gurney et al., 1994) generated hSOD1G93A-UeGFP mice, which allowed detailed study of CSMN health. We observed a progressive degeneration of eGFP+ CSMN, as previously reported (Ozdinler et al., 2011), with apical dendrite vacuolation and presence of autophagosomes, suggesting an ongoing intrinsic cellular degeneration. This novel reporter mouse model for ALS, now allows the detailed cellular analysis of CSMN without the requirement of retrograde labeling surgeries, which was not possible before.

The main challenge of genetic labeling approaches remains identification of genes/promoters that distinctly and specifically labels a neuronal subpopulation of interest. Once such reporters have been identified and characterized, there are numerous advanced applications. Recent advances in microscopy allow imaging cells in live animals (intravital imaging) (Pittet and Weissleder, 2011; Weigert et al., 2013). *In vivo* imaging has recently been applied to study spinal cord regeneration (Laskowski and Bradke, 2013). Optogenetics approaches take advantage of channelrhodopsin-2, which can be genetically targeted to neurons of interest and used to manipulate neural activity with millisecond precision using light (Packer et al., 2013). This allows imaging and long-range circuit mapping of multiple cell types (Atasoy et al., 2008). Optophysiology also grants success in such studies (Smedemark-Margulies and Trapani, 2013). Recently, optogenetic reactivation of hippocampal neurons activated during fear conditioning was shown to activate fear memory recall induced by light (Liu et al., 2012). Mosaic analysis with double markers (MADM) uses a site-specific recombinase to catalyze inter-chromosomal recombination that creates sparsely labeled neurons only after recombination (Luo, 2007). Type, density, and timing of labeling can be controlled by tissue-specific expression of the recombinase. Genetically encoded fluorescent protein-based biosensors can be used to study a broad assortment of signaling molecules and networks (Depry et al., 2013). For example, fluorescent proteins can be used to monitor proteinprotein interactions in living cells using fluorescent resonance energy transfer (FRET) microscopy (Stepanenko et al., 2011; Day and Davidson, 2012). A recent nanobody-based technology utilizes fluorescent proteins commonly used to label cells and neurons as scaffolds for cell-specific gene manipulation allowing selective modulation of diverse GFP-labeled cells across transgenic lines such as optogenetic probing of neural circuits (Tang et al., 2013).

Numerous techniques and applications are being developed simultaneously to target neurons of interest. Advances in genetic labeling of select sets of neuron populations coupled with numerous applications that allow their cellular and molecular analysis at different experimental conditions and disease settings will reveal the underlying mechanisms for proper cellular function as well as the molecular networks that are responsible for their selective vulnerability. This information will be the foundation for future effective treatments strategies in diseases that primarily affect distinct neuron populations at initial stages.

#### **CONCLUSION**

Techniques allow detailed analysis, but it is our critical thinking that shapes the field and the future. Many important discoveries are made when approaches and critical thinking develop simultaneously. The definition of "disease," the description of pathologies, and our understanding of symptom development have vastly changed in the last decade. In addition, numerous new techniques and applications, which allow studies that were not previously possible, have become available. These two critical components of innovation have the potential to shape the future of neurodegenerative diseases once understood and applied properly. We now have reasons to believe that numerous discoveries will emerge in the near future.

Recently, the word "spectrum" was introduced when describing numerous neurodegenerative disorders to highlight the blurred boundaries among diseases and place them under an umbrella of systems degeneration with many common features. This is an important shift from disease-based thinking toward a more mechanism-oriented understanding that emphasizes the common underlying pathologies that give rise to different forms of neurodegeneration. This paradigm shift in our understanding has an impact on defining pathologies, disease mechanisms, and symptoms. The neurons that show vulnerability and undergo progressive degeneration have moved center stage. We now realize the importance of understanding the cellular, molecular, and genetic basis of pathology at a cellular level. This change in critical thinking proved to be correct by the failures of clinical trials. The expectation for improved lifespan in mice to translate into success in ALS patient survival is now considered mostly unrealistic.

**FIGURE 3 | Genetic Labeling of CSMN. (A)** Schematic drawing of genetic labeling of CSMN by eGFP expression under control of UCHL1 promoter. **(B)** A representative sagittal section of a P120 mouse brain showing eGFP+ CSMN and their axons. **(C)** In motor cortex, eGFP+ CSMN can be seen in

layer 5a and 5b. **(C')** Higher magnification of individual representative eGFP+ CSMN in the motor cortex. **(D–F)** eGFP+ axons of CSMN projecting through the striatum **(D)**, pons **(E)**, and pyramidal decussation **(F)**. Scale bars: **(B)** = 1 mm, **(C,D** and **E)** = 200μm, **(C')** = 20μm, **(F)** = 500μm.

Today, clinical companies are more cautious when making a decision to move forward, and seek more cell-based evidence that supports motor neuron survival (Genç and Ozdinler, 2013).

In recent years, there have been significant changes in the meaning of the word "symptom." Previously, symptoms were described as signs of pathology that can be detected by visual examination or various tests. However, it became obvious that our ability to detect signs of pathology at a molecular, cellular, and systems level was different. Even when no obvious signs were present, the pathology was taking its course at a cellular level. If the cellular basis of disease causing pathologies was understood, then these findings would set the stage, not only for early detection of the disease, but also for the development and implementation of effective long-term treatment strategies. Therefore, it is important to detect symptoms at a cellular level before pathologies become evident. We need to develop new approaches to focus our attention on the neuron, on the cell that is affected and has become vulnerable. Because if we understand the molecular and genetic basis of neuronal vulnerability at a cellular level, then we will have the chance to understand the basis of neuronal pathology and systems degeneration.

Studying CSMN biology is especially challenging, not only because they are limited in numbers and not easy to identify, but also because the techniques, technologies, approaches, and model systems to study them are not easy to develop. Due to species differences, modeling CSMN degeneration in mice has intrinsic limitations. Even though a minor defect would lead to a motor dysfunction in patients, mice would not display an obvious defect. This could indeed be one of the reasons why the mouse models of motor neuron diseases do not show a prominent phenotype. Working in models without an obvious functional readout can be challenging. However, if the models are built at a cellular level, or if the focus is shifted from mice to the motor neurons in mice, then the picture and the scope of perception changes dramatically. At a cellular level, the CSMN in humans and in mice are almost identical. It is thus important to develop technologies that reveal the underlying factors that contribute to the cellular vulnerability and degeneration. As long as our focus is the neuron and not the mice, the information gathered from healthy and diseased neurons would be translational.

Understanding the controls over CSMN health is becoming very central to numerous neurodegenerative diseases. Here we reviewed the exciting progress in the study of upper motor neurons. Retrograde labeling is a prominent approach to identify the location of the neuron and to visualize its cell body. AAVmediated retrograde transduction, on the other hand, is not only a powerful tool to deliver genes of interest, but also important to reveal details of cellular cytoarchitecture. However, both of these applications require surgery and the success of each experiment mostly depends on surgical expertise and mouse survival. Genetic labeling that intrinsically targets a distinct neuron population offers a solution for the potential limitations of retrograde labeling surgeries. Generation and characterization of the UCHL1-eGFP mouse, in which the CSMN are genetically labeled, has been pivotal for overcoming numerous important limitations by allowing *in vivo* visualization and cellular analysis of neurons that are vulnerable in neurodegenerative diseases.

With the advancements made in the study of CSMN biology, the future awaits numerous important discoveries. FACS purification coupled with *in vitro* culturing will allow novel drug screening and verification platforms using motor neuron health as a readout for success. This application has the potential to discover new molecules and compounds for future clinical trials. In addition, development of AAV-mediated gene delivery, together with the identification of new genes and pathways that are important for disease pathology will enable direct cellular therapies into the motor cortex of patients improving the survival of upper motor neurons and enhancing their connectivity. Identification of the cellular and molecular basis of CSMN vulnerability will be revealed, paving the way for understanding the cellular pathways, networks, and dynamics that are important to improve motor neuron health.

We live in very exciting times due to the numerous improvements in our critical thinking of neurodegenerative diseases, and the technologies that support discoveries are developing faster than ever. The new knowledge gathered from neurons of interest will enable development of novel effective long-term treatment strategies and has the potential to expedite new discoveries that will enable improved health in patients.

#### **REFERENCES**

ER-associated pathway. *J. Neurochem.* 114, 795–809. doi: 10.1111/j.1471- 4159.2010.06806.x

spinal motor neurons in an ALS mouse model. *J. Neurosci.* 33, 7890–7904. doi: 10.1523/JNEUROSCI.2787-12.2013

**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.

*Received: 30 November 2013; accepted: 10 March 2014; published online: 26 March 2014.*

*Citation: Jara JH, Genç B, Klessner JL and Özdinler PH (2014) Retrograde labeling, transduction, and genetic targeting allow cellular analysis of corticospinal motor neurons: implications in health and disease. Front. Neuroanat. 8:16. doi: 10.3389/fnana. 2014.00016*

*This article was submitted to the journal Frontiers in Neuroanatomy.*

*Copyright © 2014 Jara, Genç, Klessner and Özdinler. 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) or licensor 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.*

## Evolutionary development of the amygdaloid complex

#### *Mohan Pabba\**

*Neurosciences Unit, Department of Cellular and Molecular Medicine, Faculty of Medicine, University of Ottawa, Ottawa, ON, Canada \*Correspondence<EMAIL_ADDRESS><EMAIL_ADDRESS>
#### *Edited by:*

*Makoto Fukuda, Baylor College of Medicine, USA*

#### *Reviewed by:*

*Joshua Corbin, Children's National Medical Center, USA*

**Keywords: amygdala, anatomy, tetrapods, mammals, evolution**

In the early 19th century, Burdach discovered an almond-shaped mass of gray matter in the anterior portion of the mammalian temporal lobe, which he called "amygdala" (Burdach, 1819–1822). The first anatomical description of the amygdala was made in 1867 by Meynert (1867). Subsequently, a large number of other nuclei were added to the amygdala to constitute what is now known as the "amygdaloid complex" (AC) (Johnston, 1923). Until this day, AC remains a subject of intense investigation in terms of content and evolutionary development since it is a much more complicated structure than what was previously thought. It is therefore, important to know the evolutionary developmental origin of AC before we can completely understand its function.

The AC is a multinuclear complex comprised of 13 nuclei. These nuclei are divided into three major groups: the basolateral, cortical-like, and centromedial. Other accessory nuclei such as the intercalated cell masses (I) and the amygdalo-hippocampus area have also been described. The basolateral group is comprised of the lateral nucleus (LA), basal nucleus (B), and accessory basal nucleus (AB) (Johnston, 1923). The cortical-like group of nuclei includes the nucleus of the lateral olfactory tract (NLOT), bed nucleus of the accessory olfactory tract (BAOT), anterior and posterior cortical nuclei (CoA and CoP, respectively), and periamygdaloid cortex (PAC). The centromedial nucleus consists of the central nucleus (CeA), medial nucleus (M), and amygdaloid part of the bed nucleus of stria terminalis (BST). The major remaining groups of AC are the amygdalohippocampal area (AHA) and intercalated nuclei (I) (Aggleton, 2000; Sah et al., 2003). These different nuclei of AC are connected within and also with various brain regions, and thus, process various types of information (e.g., olfactory and **Figure 1A**).

Swanson and Petrovich (1998) definition of amygdala as neither a structural nor a functional unit provides an attractive point to explore the evolutionary developmental aspects of AC because a growing number of evidence suggests AC as an evolutionarily conserved structure. Earlier, research on structural organization of AC in different amniotic vertebrates revealed a common pattern of organization, along with shared functional roles. Conversely, research on anamniotes provided little comparative information regarding structural organization of AC. However, recent studies have shown a homology between amygdaloid components of amniotes and anamniotes. To better understand the evolutionary and developmental history of a particular brain region, one needs to follow a "sequential (step by step) approach," which takes into account the developmental, topological, hodological, genetical, and functional history. Interestingly, recent data on AC of mammals, reptiles, and anurans suggest that the evolution of AC occurred as common traits of telencephlon, for example, regions of cortical amygdala such as nLOT and accessory olfactory bulb (AOB) (Remedios et al., 2007; Huilgol et al., 2013); but not as the sum of unrelated structures with different origins. The present understanding of AC in developmental and adult vertebrates suggests two major divisions of telencephlon: the pallium and the subpallium (Puelles et al., 2000; Martinez-Garcia et al., 2002; Moreno and Gonzalez, 2007b; Remedios et al., 2007). This dual view or origin makes AC a histogenetic complex structure of the adult brain, with extremely intense morphogenetic and migratory processes during the development in all tetrapods (Puelles et al., 2000). In mammals, the pallial component is composed of "cortical amygdala" and "basolateral amygdala." In turn, the subpallial component consists of the striatal component, central amygdala, and medial amygdala. This basic plan is shared by reptiles, birds, and also by anuran amphibians (Martinez-Garcia et al., 2002; Medina et al., 2004). Interestingly, this basic description is possible only in few mammalian tetrapods, but not in the non-mammalian amniotes and the anurans where they have no clear anatomical subdivisions. The existence of shared embryological AC components in all tetrapods provides clues to the presence of precursors of the amygdaloid nuclei from anamniotes (Moreno and Gonzalez, 2007b). The following sections deal with the current view on accepted and shared components of AC in tetrapods.

The amygdala is a part of a phylogentically conserved olfactory system, particularly the olfactory bulb, in vertebrate evolution in terms of embryological origin, neurochemistry, connectivity, and function (Martinez-Garcia et al., 2002; Huilgol et al., 2013). Additionally, a major part of amygdala is also an integral component of the vomeronasal system of the tetrapod (except avian) brain (Swanson and Petrovich, 1998; Moreno and Gonzalez, 2005a).

In mammals, the vomeronasal information passes via the AOB to medial (MeA) and cortical postero-medial amygdala (CoApm) (Swanson and Petrovich, 1998). In addition, the amygdala also receives information from the main olfactory bulb (MOB) and hypothalamus to modulate reproductive and defensive behaviors (Canteras et al., 1995). In reptiles and anurans, the existence of a well-developed "vomeronasal amygdala"

has also been reported (Moreno and Gonzalez, 2003), although no vomeronasal amygdaloid nuclei has been described in birds so far (Martinez-Garcia et al., 2006). Thus, in all tetrapods, the main secondary vomeronasal brain areas belong to AC.

changes in the position of certain AC nuclei during the course of

In mammals, the olfactory amygdaloid system consists of the distinct cortical (CoA, CoP nuclei, etc.) and basolateral amygdala (BL, M, LA nuclei). LA receives major sensory input, and is important for emotional behavior (Ledoux et al., 1990). Studies on birds indicate the presence of nuclei that are comparable to CoP of the mammalian amygdala. These studies also revealed the possession of counterparts similar to BM and LA of the mammalian amygdala (Martinez-Garcia et al., 2006). In reptiles, studies on the olfactory system showed comparable functional circuitry with the mammalian BA complex. The anuran counterpart of the mammalian olfactory amygdala is LA (Moreno and Gonzalez, 2004). This part of the amygdala in anurans receives directly or indirectly olfactory, visual, auditory, somatosensory, vomeronasal, and gustatory information. The observed integration that occurs in AC of tetrapods is responsible for the acquisition of "emotional memory," which pertains to the survival of individuals during their defence against danger, their

and Gonzalez (2007b).

interaction with sexual partners, or their fight with an enemy (Ledoux, 2000). Therefore, the amygdala receives large sensory input information from olfactory and vomeronasal projections, and are conserved in tetrapods.

Studies on tetrapods showed CeA as the main receiver of a wide range of sensory information from other amygdaloid regions in addition to the thalamus and brain stem. Moreover, CeA is known to link and integrate the emotional and motor components of behavior (Han et al., 1997), behavioral responses to nociceptive and visceral pain (Han and Neugebauer, 2004), and behavioral responses to stressful stimuli (Saha et al., 2000). CeA also mediates many of the autonomic, somatic, endocrine, and behavioral responses in different tetrapods. The autonomic amygdala provides a link between environmental stimuli and animal behavioral responses, and thus, provides an important significance in terms of evolutionary conservation.

Another conserved shared system of tetrapods is the strong amygdalohypothalamic connections (Martinez-Marcos et al., 1999; Moreno and Gonzalez, 2005a). In mammals, nuclei that project to hypothalamus through the stria terminalis arise from the medial and basolateral amygdala (Swanson and Petrovich, 1998). As in mammals, the amygdalohypothalamic projections of anurans (Moreno and Gonzalez, 2005b) also project through the stria terminalis. In anurans, the amygdalo-hypothalamic connections control functions mediated by the hypothalamus in response to pheromones and odors (Reiner and Karten, 1985; Swanson and Petrovich, 1998).

As in mammals, the amygdalohypothalamic projections of anurans, project through the stria terminalis. The main similarity with amniotes is the projection to the hypothalamus from comparable amygdaloid territories carrying vomeronasal, olfactory, and multimodal information (Reiner and Karten, 1985; Swanson and Petrovich, 1998). The situation of amygdalo-hypothalamic projections in birds is more complicated because of the lack of a well-developed olfactory/vomeronasal system.

On the other hand, studies that compare the distribution of neuronal markers (either proteins or genes/transcription factors) across the development of analogous AC nuclei from different species as well as within the same species have also provided valuable information on the evolution of AC. For instance, similarities in the molecular profiles of the pallium and subpallium of mice and chickens were obtained by comparing the nested expression domains of genes such as Dlx-2, Tbr-1, Pax-6, NKx-2.1, and Emx-1 (Puelles et al., 2000). Moreno and Gonzalez, using the distribution of somatostatin, nitric oxide synthase, etc. in anuran CeA and MeA in a series of studies, has predicted that these

parts of AC could be related to AC of amniotes (Moreno and Gonzalez, 2005a, 2007a). Medina et al. demonstrated a possible existence of evolutionary relationship in various AC nuclei of mammals, reptiles and birds by testing the expression patterns of genes/transcription factors such as Lhx2, Lhx9, Pax6, Islet 1, NKx2, Lhx6, and Lhx5 in forebrain regions of these animals (Medina et al., 2011). Using the similarities and differences in the expression of Lhx1 and Lhx5, Abellan et al. suggested a common pattern of evolutionary conservation in telencephalon between mice and chickens during various stages of development (Abellan et al., 2010). By examining the distribution of Lhx2, Trb1, reelin, and CdK5, Remedios et al. estimated a possible developmental and evolutionary link between nLOT of AC and neocortex (Remedios et al., 2007; Subramanian et al., 2009). Another study from Tole's group, using migratory genetic markers (e.g., NP2 and AP2α), showed that the distinct halves of AOB [posterior and anterior (pAOB and aAOB)] has different developmental origins, and that pAOB could be a component of AC as it was positioned closely to MeA in anamniotes (*Xenopus*) (Huilgol et al., 2013). Finally, Barger et al. using a different approach, i.e., by comparing the percentage of neurons in individual nuclei of AC between humans and apes, suggested that during the course of human evolution, LA of AC has further progressed in humans (Barger et al., 2012).

#### **CONCLUDING REMARKS**

The classical hypothesis proposed by Edinger regarding the evolution of the brain attracted much attention (Edinger, 1908). He proposed that the telencephlon evolved in progressive stages of complexity and size, culminating to the human brain. He also stated that there is an "old brain" (the subpallium at the telencephalic base) followed by the addition of a "new brain" (the pallium at the top of the telencephlon). Nevertheless, this classical hypothesis provides evidence on the existence of a basic plan in the origin, regionalization, and organization of the forebrain of vertebrates. Based on the data pertaining to the organization of AC, there seems to have several important features that are common to all tetrapods: (1) it is formed by pallial and subpallial derivatives; (2) it is topographically situated in the ventrolateral caudal telencephalic hemispheres; (3) it has shared features in relation with different functional systems like the vomeronasal, olfactory, autonomic, and multimodal systems along with an intricate intra-amygdaloid network; (4) it is the origin of important hypothalamic projections; (5) it has a common embryological origin for several prominent features of AC; (6) it has the presence of a main output for autonomic system; finally, (7) it has abundant local circuit neurons that are shared by most amniotes. Thus, in the light of recent findings on AC (Remedios et al., 2007; Butler et al., 2011; Huilgol et al., 2013) also strongly support the idea that tetrapods share the same basic plan.

The increase in size of the pallium, especially in mammals, has an evolutionary importance. The current spatial arrangements of the mammalian AC are still found in living anurans. Therefore, it is now obvious that these "new evolutionary nuclei" would have pushed the "most conserved nuclei" (Moreno and Gonzalez, 2007b). It explains why mammals have the central, medial, and basolateral nuclei occupying the most medial positions, whereas the cortical amygdaloid nuclei occupy the most lateral positions (**Figure 1B**) (Moreno and Gonzalez, 2007b). Consequently, the brain of ancestral tetrapods developed an elaborate AC in response to new requirements imposed on them as a part of the transition from water-to-land. Therefore, the basic organization of the brain system, at least in the case of the AC, is still recognizable in all existing tetrapods, and can be compared with that of mammals.

#### **ACKNOWLEDGMENTS**

I am thankful to Dr. Richard Bergeron, Dr. Marzia Martina, and Wissam B. Nassrallah for their helpful feedback on the manuscript.

#### **REFERENCES**

et al. (2012). Neuronal populations in the basolateral nuclei of the amygdala are differentially increased in humans compared with apes: a stereological study. *J. Comp. Neurol.* 520, 3035–3054. doi: 10.1002/cne.23118

transition. *J. Anat.* 211, 151–163. doi: 10.1111/j.1469-7580.2007.00780.x

*Received: 09 July 2013; accepted: 06 August 2013; published online: 28 August 2013.*

*Citation: Pabba M (2013) Evolutionary development of the amygdaloid complex. Front. Neuroanat. 7:27. doi: 10.3389/fnana.2013.00027*

*This article was submitted to the journal Frontiers in Neuroanatomy.*

*Copyright © 2013 Pabba. 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) or licensor 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.*

## Investigation of spinal cerebrospinal fluid-contacting neurons expressing PKD2L1: evidence for a conserved system from fish to primates

*Lydia Djenoune1,2,3,4,5,6, Hanen Khabou1,2,3,4‡, Fanny Joubert 4,7‡, Feng B. Quan5,6, Sophie Nunes Figueiredo1,2,3,4, Laurence Bodineau4,7, Filippo Del Bene8,9,10, Céline Burcklé1,2,3,4†, Hervé Tostivint 5,6 and Claire Wyart 1,2,3,4\**

*<sup>1</sup> Institut du Cerveau et de la Moelle Épinière, Hôpital de la Pitié-Salpêtrière, Paris, France*

*<sup>5</sup> Muséum National d'Histoire Naturelle, Paris, France*

*<sup>8</sup> Institut Curie, Paris, France*

*<sup>9</sup> Centre National de la Recherche Scientifique UMR 3215, Paris, France*

*<sup>10</sup> Institut National de la Santé et de la Recherche Médicale U 934, Paris, France*

#### *Edited by:*

*Laurent Gautron, The University of Texas Southwestern Medical Center, USA*

#### *Reviewed by:*

*Marina Bentivoglio, Université di Verona, Italy Ramon Anadon, Universidad de Santiago de Compostela, Spain*

#### *\*Correspondence:*

*Claire Wyart, Institut du Cerveau et de la Moelle épinière, Hôpital de la Pitié-Salpêtrière, 83, bld de l'hôpital, 75013 Paris, France e-mail<EMAIL_ADDRESS>
#### *†Present address:*

*Céline Burcklé, Institut de Biologie du Développement de Marseille, Marseille, France*

*‡These authors have contributed equally to this work.*

Over 90 years ago, Kolmer and Agduhr identified spinal cerebrospinal fluid-contacting neurons (CSF-cNs) based on their morphology and location within the spinal cord. In more than 200 vertebrate species, they observed ciliated neurons around the central canal that extended a brush of microvilli into the cerebrospinal fluid (CSF). Although their morphology is suggestive of a primitive sensory cell, their function within the vertebrate spinal cord remains unknown. The identification of specific molecular markers for these neurons in vertebrates would benefit the investigation of their physiological roles. PKD2L1, a transient receptor potential channel that could play a role as a sensory receptor, has been found in cells contacting the central canal in mouse. In this study, we demonstrate that PKD2L1 is a specific marker for CSF-cNs in the spinal cord of mouse (*Mus musculus*), macaque (*Macaca fascicularis*) and zebrafish (*Danio rerio*). In these species, the somata of spinal PKD2L1+ CSF-cNs were located below or within the ependymal layer and extended an apical bulbous extension into the central canal. We found GABAergic PKD2L1-expressing CSF-cNs in all three species. We took advantage of the zebrafish embryo for its transparency and rapid development to identify the progenitor domains from which *pkd2l1*+ CSF-cNs originate. *pkd2l1*+ CSF-cNs were all GABAergic and organized in two rows—one ventral and one dorsal to the central canal. Their location and marker expression is consistent with previously described Kolmer–Agduhr cells. Accordingly, *pkd2l1*+ CSF-cNs were derived from the progenitor domains p3 and pMN defined by the expression of *nkx2.2a* and *olig2* transcription factors, respectively. Altogether our results suggest that a system of CSF-cNs expressing the PKD2L1 channel is conserved in the spinal cord across bony vertebrate species.

**Keywords: cerebrospinal fluid-contacting neurons (CSF-cNs), PKD2L1, mouse, macaque, zebrafish, GABAergic neurons, spinal cord**

**Abbreviations:** cc, Central canal; CSF, Cerebrospinal fluid; CSF-cNs, Cerebrospinal fluid-contacting neurons; dpf, days post fertilization; FISH, Fluorescent *in situ* hybridization; GAD, Glutamic Acid Decarboxylase; GABA, γ-aminobutyric acid; GFP, Green Fluorescent Protein; GFP+, GFP positive; hpf, hours post fertilization; IHC, Immunohistochemistry; KAs, Kolmer–Agduhr cells; MAP2, microtubule-associated protein 2; PB, Phosphate Buffer; PBS, Phosphate Buffered Saline; PFA, paraformaldehyde; *pkd2l1*, mRNA form of polycystic kidney disease 2-like 1 in zebrafish; *pkd2l1*+, *pkd2l1* positive; PKD2L1, polycystic kidney disease 2-like 1 protein in mouse and macaque; PKD2L1+, PKD2L1 positive; TRP, Transient Receptor Potential; VGAT, vesicular GABA transporter.

#### **INTRODUCTION**

Spinal cerebrospinal fluid-contacting neurons (CSF-cNs) were identified and described by Kolmer (1921, 1931) and Agduhr (1922) in over 200 species based on cellular morphology, location, and Nissl staining. They independently noted that these cells exhibit an apical bulbous extension in the central canal and send basal axonal projections to other cells. Their observations suggested that CSF-cNs could constitute a sensory organ (referred to as the sagittal organ by Kolmer) interfacing the cerebrospinal fluid (CSF) with the nervous system at the level of the spinal cord.

Since the discovery of CSF-cNs, electron microscopy studies have shown that these cells exhibit a sensory tuft, previously referred to as a "brush border" (Dale et al., 1987b), a "central body" (Vigh and Vigh-Teichmann, 1973) or a "bud" (Stoeckel et al., 2003). The apical bulbous extension projects from the perikarya of CSF-cNs toward the central canal and ends with a terminal bud in contact with its lumen. This apical extension is characterized by the dendritic marker microtubule-associated protein 2 (MAP2) (Orts-Del'Immagine et al., 2014).

 In African clawed frog and zebrafish, these GABAergic CSF-cNs were named Kolmer–Agduhr cells (KAs) to distinguish them from ciliated ependymal cells (Dale et al., 1987a; Bernhardt et al., 1992). In these species CSF-cNs project an ascending axon ventrally in the spinal cord (Dale et al., 1987a; Bernhardt et al., 1992; Higashijima et al., 2004a; Wyart et al., 2009).

Despite the anatomical studies of CSF-cNs and their implication in modulating locomotion (Wyart et al., 2009), their physiological role in vertebrates remains unknown. One obstacle to answering this question is the lack of a specific genetic marker to identify these cells. Recently the calcium-permeable polycystic kidney disease 2-like 1 (PKD2L1) channel (Huang et al., 2006; Ishimaru et al., 2006), first identified in kidney, retina, and heart (Basora et al., 2002) was found to be expressed in cells contacting the CSF at the level of the brainstem and spinal cord in mouse (Huang et al., 2006; Orts-Del'Immagine et al., 2012). This channel belongs to the family of Transient Receptor Potential (TRP) channels. These are known to be chemo-, thermo- or mechanosensitive (Delmas, 2004; Delmas et al., 2004). Expression of this channel and location of these cells at the interface with the CSF are consistent with the hypothesis that CSF-cNs have a proprioceptive sensory function.

In this study, we investigated whether PKD2L1 was a specific marker for CSF-cNs across vertebrates, by examining mouse, macaque and zebrafish. In mouse and macaque PKD2L1 was enriched in the sensory tuft of CSF-cNs in cervical, thoracic and lumbar spinal cord. The soma of PDK2L1+ cells was located in the ependymal layer or just underneath. In both species, a significant proportion of PKD2L1+ CSF-cNs was distinctly identified as GABAergic. We took advantage of the zebrafish model organism to thoroughly characterize the properties and lineage of *pkd2l1*+ cells in the spinal cord. In this model, all *pkd2l1*+ cells contacting the central canal were confirmed as GABAergic neurons. Reciprocally, all KAs, defined by their location and their GABAergic phenotype expressed *pkd2l1*. Using specific transgenic lines, we confirmed that these cells derive from two progenitor domains. The dorsal *pkd2l1*+ KAs originated from pMN and were labeled by the Enhanced Green Fluorescent Protein (EGFP) in the *Tg(olig2:EGFP)* while the ventral *pkd2l1*+ KAs originated from p3 and were labeled by *Tg(nkx2.2a:mEGFP)*. Altogether our data show that PKD2L1 is a specific marker of CSF-cNs shared between multiple vertebrate species. We found evidence for PKD2L1-expressing GABAergic CSF-cNs in all examined species. Our results on the developmental origin of CSF-cNs in zebrafish open new paths of investigation in mammals.

#### **MATERIALS AND METHODS**

#### **MOUSE**

#### *Experimental animals*

Wild type (WT) mice (*Mus musculus*, OF1 strain) were kept on a 12 h light/dark cycle with free access to food and water. Experiments were performed at embryonic (E), postnatal (P) and adult stages: four WT adult mice, one WT newborn, and three WT embryos were used in the present study. One adult male and two E16.5 GAD67-GFP knock-in embryos (Tamamaki et al., 2003), resulting of the insertion of the cDNA encoding EGFP into the locus encoding GAD67, have been used. After mating, the day of detection of the vaginal plug was considered as embryonic day E0.5. Pregnant females were sacrificed by decapitation at E14.5, E16.5, or E18.5. Uterine horns were removed from the mother and embryos were excised from their individual bag, soaked in cold 0.1 M Phosphate Buffer (PB) for 10 min and fixed by immersion in cold 2% paraformaldehyde (PFA) solution in 0.1 M PB for 3 h at 4◦C. Newborn mice (P1) were deeply anesthetized by an exposure to low temperature and the spinal cord was immediately removed and fixed with 2% PFA for 4 h. Adult mice were deeply anesthetized with intraperitoneal injection of Nembutal® (150 mg·kg−1) and transcardially perfused sequentially with NaCl 0.9% (20 ml) and PFA (2%, 30 ml for WT adults and 4%, 30 ml for the GAD67-GFP mouse). After fixation, spinal cords were dissected out and post-fixed for 24 h in the fixative solution (2% PFA for WT mice and 4% for the GAD67-GFP mouse) at 4◦C. Following fixation (and post-fixation if appropriate), tissues were cryo-protected for 24–48 h in 30% sucrose in 0.1 M PB at 4◦C and then stored at -20◦C for later use. Experiments were performed in accordance with the European Communities Council (EEC) Directive of 22 September 2010 (2010/63/EU) and French law (87/848).

#### *Immunohistochemistry*

Standard immunofluorescence procedures were used to localize antigens of interest (Voituron et al., 2011). At the cervical level, 10μm-thick coronal sections of the spinal cord were obtained using a cryostat (Leica CM 3050S), mounted on silanized slides and stored at −20◦C for later IHC using a rabbit anti-Polycystin-L (anti-human PKD2L1, 1:500 dilution; Millipore, Billerica, MA, USA). This PKD2L1 antibody was raised against the synthetic peptide from the N-terminal of human Polycystin-L. For PKD2L1 IHC at E14.5 and E16.5, we used an Antigen Unmasking Solution (Vector Laboratories). Briefly, the sections were incubated in 1% Bovine serum albumin (BSA) and 0.1% Triton X-100 for 20 min at room temperature (RT). After rinsing with 0.1 M phosphate buffered saline (PBS), the sections were incubated for 30 min at RT with DAPI (1.25μg/ml; Life Technologies) and with Alexaconjugated anti-rabbit secondary antibodies (1:500 dilution; Life Technologies). For GFP and PKD2L1 double IHC, 30μm-thick coronal floating sections obtained using a cryostat (Leica CM 1850S) were immunostained by co-incubating two primary antibodies, a chicken anti-GFP (1:1000 dilution; Abcam, Cambridge, UK) with the rabbit anti-PKD2L1 (1:500 dilution) overnight at 4◦C, and then the specific secondary antibodies Alexa Fluor 488 donkey anti-chicken IgG (1:500 dilution, Life Technologies) and Alexa 555-conjugated goat anti rabbit for 2 h at RT. In all IHC experiments, control sections were processed in parallel by omitting primary antibodies. Finally, sections were washed with PBS before being mounted onto glass slides with fluorescent mounting medium (AquaPolyMount, Biovalley, Marne La Vallée, France). Sections were observed under an Olympus FV1000 confocal microscope.

#### *Quantification of cells*

On 10μm-thick sections, we performed Z-stacks with a step size of 1–1.5μm in a region of interest (ROI) centered on the central canal (approximately 150 × 220μm width, *n* = 13 sections for E16.5 GAD67-GFP and 115 × 140μm width, *n* = 7 sections for adult GAD67-GFP sections). For each analyzed section, cells were identified by the DAPI staining in order to avoid counting multiple times the same cell. Within the ROI, all PKD2L1+ cells were counted manually and probed for GFP in the GAD67-GFP line.

#### **MACAQUE**

#### *Experimental animals*

Tissue from two adult macaques (*Macaca fascicularis*, a 25-year old female and an 8-year old male) was used for this study. These animals were sacrificed for other purposes and spinal cord tissue samples were subsequently collected and devoted to the present study. Spinal cord collection was performed after the sacrifice of animals, in strict accordance with the recommendations of the Weatherall Report regarding good animal practice. All surgical procedures and experimental protocols were carried out in strict accordance with the National Institutes of Health guidelines (2013) and the recommendations of the EEC (2010/63/EU).

#### *Tissue extraction and preparation*

Macaques were deeply anesthetized with intramuscular injection of ketamine and xylazine at 50 and 5 mg/kg, respectively. Following the anesthesia, each animal was exsanguinated and perfused with 0.9% NaCl before their tissues were fixed by an intracardial perfusion containing 4% PFA in 0.1 M PBS. Spinal cord was extracted and meninges were removed using fine forceps. Tissue was embedded in 5% agarose and sectioned at 50μm using a vibratome (Leica VT1000S). Additionally, frozen tissue blocks were obtained by passing tissue through a series of cryoprotection solutions (20, 30% sucrose in PBS) and freezing in isopentane; frozen sections were cut at 20μm (HM 650V Microtome, Thermo Scientific). All sections were stored in 0.4% Sodium Azide in 0.1 M PBS at 4◦C until IHC.

#### *Immunohistochemistry*

Free-floating sections from cervical, thoracic, and lumbar spinal cord tissue were used for IHC. Sections were first incubated in a blocking solution composed of 1% BSA, 0.5% Triton X-100 and 2% normal goat serum in PBS to reduce non-specific labeling. Sections were then incubated in primary antibodies overnight at 4◦C. After multiple washes, sections were incubated in secondary antibodies for 4 h at 4◦C.  Secondary antibodies were Alexa Fluor 488- or Alexa Fluor 568-conjugated anti-rabbit (1:500; Life Technologies) and Alexa Fluor 488-antiguinea pig (1:500; Life Technologies). The same blocking solution as previously was used to dilute the antibodies. PBS was used for washing sections between each step. Sections were mounted on glass slides, cover slipped with Prolong® Gold Antifade Reagent (Life Technologies) mounting medium and then imaged on a Leica SP2 AOBS AOTF inverted confocal laser scanning microscope. Single optical slice images of individual labeled cells were taken using a 63× oil objective.

#### **ZEBRAFISH**

#### *Experimental animals*

All zebrafish (*Danio rerio*) lines were maintained and raised on a 14/10 h light cycle and water was regulated at 28.5◦C, conductivity at 500μS and pH at 7.4 (Westerfield, 2000). WT AB and Tüpfel long fin (TL) embryos were used for whole mount *in situ* hybridization (ISH). *Tg(nkx2.2a:mEGFP)* (Ng et al., 2005; Kirby et al., 2006) and *Tg(olig2:EGFP)* (Shin et al., 2003) transgenic lines were kindly provided by Prof. Bruce Appel, University of Colorado, Denver, USA. Embryos were dechorionated and staged according to number of somites as described (Kimmel et al., 1995): the 30-somite stage corresponds to Prim-5 or 24 h post fertilization when raised at 28.5◦C. Adult fish were anesthetized in 0.02% MS 222 (Sandoz, Levallois-Perret, France) and killed by decapitation. All procedures were approved by the Institutional Ethics Committee at the Institut du Cerveau et de la Moelle épinière (ICM), Paris, France, the Ethical Committee Charles Darwin and received subsequent approval from the EEC (2010/63/EU).

#### *Generation of the pkd2l1 probe*

To generate the *pkd2l1* ISH probe, the coding fragment for *pkd2l1* was amplified from zebrafish embryo total cDNA using the following primers (5 to 3 ): pkd2l1\_For: TAGTGGTGATACTGCTTGCTGTGGTGG (from the end of the exon 6 of the *pkd2l1* gene and the beginning of exon 7) and pkd2l1\_Rev: TGGTTCCACACTGTTCTCGAGGTCACG (from the end of exon 13). The PCR fragment was cloned into the pCRII-TOPO vector (Life Technologies, Carlsbad, CA, USA). The resulting plasmid was linearized with NotI. The *gad67* plasmid, kindly provided by Dr. Uwe Strähle, Karlsruhe Institute of Technology, Germany, was linearized with NcoI. Digoxigenin (DIG)- and fluorescein (Fluo)-labeled probes were synthesized using SP6 RNA polymerase with the RNA Labeling Kit (Roche Applied Science, Basel, Switzerland) to generate both *pkd2l1* and *gad67* antisense probes. To generate *pkd2l1* sense probes, the plasmid was linearized with KpnI and transcription was carried out using T7 RNA polymerase. All probes were purified using the mini Quick Spin RNA Column (Roche, Basel, Switzerland).

#### *In situ hybridization*

Whole-mount ISH were performed as previously described (Parmentier et al., 2011; Alunni et al., 2013) on embryos or dissected adult spinal cords fixed in 4% PFA in PBS overnight at 4◦C. To reveal *pkd2l1* expression, probes were detected with anti-DIG or anti-Fluo antibodies conjugated to alkaline phosphatase followed by a chromogenic reaction using a solution of NBT/BCIP as substrate (Roche Diagnostics, France). To quantify cell density or ascertain transcript colocalization, probes were detected by the antibodies conjugated to horseradish peroxidase and were revealed by Tyramide Signal Amplification using Tyramide-FITC or Tyramide-TAMRA as substrates. The specificity of the *pkd2l1* probe was verified using a *pkd2l1* sense probe as negative control (data *not shown*). Double fluorescent *in situ* hybridization (FISH) for *pkd2l1* and *gad67* were performed on adult zebrafish spinal cords and coupled with respectively DIG and Fluo.

#### *Immunohistochemistry and sectioning*

The following primary antibodies were used for IHC: rabbit anti-GABA (1:2000, Sigma-Aldrich, St. Louis, MO, USA), rabbit anti-GAD65/67 and chicken anti-GFP (both used at 1:500 dilution, Abcam, Cambridge, UK). Immunostaining specificity was established by omitting the primary specific antibodies, no immunoreactive signal was observed. Agarose sections were collected as described above. In the zebrafish embryo, we performed either 50μm-thick sagittal (for anti-GAD65/67 IHC) or transverse (for anti-GFP IHC) sections. In the adult, we performed 50μm-thick sagittal and frontal sections of previously FISH stained adult spinal cords.

#### *Combination of pkd2l1 FISH with immunohistochemistry*

*pkd2l1* FISH was performed before IHC against GFP or GAD65/67: embryos were washed and immunostained with the chicken anti-GFP antibody or the rabbit anti-GAD65/67 antibody overnight at 4◦C, and then incubated with the corresponding Alexa conjugated secondary antibodies IgG (1:500, Life Technologies) combined with DAPI (2.5μg/ml, Life Technologies).

#### *Microscopy*

Whole-mount embryos stained by NBT/BCIP were mounted in 80% glycerol. Sections of adult spinal cord were mounted in a solution of Mowiol. Embryos were imaged using a Nikon AZ100M macroscope and a Leica DM5000 B Upright microscope. Adult spinal cord sections were imaged using a Nikon AZ100M macroscope. To quantify *pkd2l1*+ and GABA+ cells, stained embryos were imaged using the Fixed Stage microscope AxioExaminer Z1 equipped with a 20× water immersion objective and a Yokogawa CSU-X1 spinning disk unit (*n* = 13 embryos for *pkd2l1* and *n* = 10 for GABA). To quantify the overlap of GFP with *pkd2l1* FISH in the *Tg(nkx2.2a:mEGFP)* and the *Tg(olig2:EGFP)* transgenic embryos, 50μm-thick sections were analyzed on an Olympus FV1000 confocal microscope equipped with a 40× water immersion objective using 405, 473, and 543 nm laser lines. Images were processed using Fiji (Schindelin et al., 2012) and Adobe Illustrator (Adobe Systems, Mountain View, CA, USA) software.

#### *Quantification of cells*

To quantify the total number of *pkd2l*1+ cells revealed by FISH per embryo, Z-stacks of the entire embryos from somite 1 to 30 were acquired. Total numbers of cells were counted per somite labeled on both sides of the midline. The boundaries for each somite were established with transmitted light. The three subtypes of *pkd2l1* expressing cells were defined according to their position relative to the central canal. We distinguished: (i) the row of cells that was ventral and in proximity to the central canal, (ii) the row that was dorsal and in proximity to the central canal and (iii) the sparse cells that were dorsal and distant from the central canal. The proportion of *pkd2l1*+ cells double-labeled for GAD65/67 in 30-somite (Prim-5) embryos was quantified based on 50μm-thick sagittal sections performed after the FISH and before GAD65/67 IHC. The proportion of *pkd2l1*+ cells doublelabeled for GFP in *Tg(nkx2.2a:mEGFP)* and *Tg(olig2:EGFP)* in 30-somite embryos was quantified based on 50μm-thick transverse sections performed after the FISH and IHC. The total number of cells is given as the mean ± Standard Error of the Mean (SEM). The same method was applied for counting of GABA+ KAs. There are four GABAergic interneuron types in the zebrafish embryo (Bernhardt et al., 1992; Higashijima et al., 2004a); KAs were identified as the only GABAergic ascending neurons with a soma located just below (KA") or above (KA') the central canal.

#### **RESULTS**

#### **PKD2L1 IS EXPRESSED IN CSF-cNs FROM EMBRYONIC STAGES TO ADULTHOOD IN THE MOUSE SPINAL CORD**

PKD2L1 was originally identified in CSF-cNs at postnatal stages P1–P4 in the mouse spinal cord (Huang et al., 2006). We investigated PKD2L1 expression in the spinal cord at the embryonic, postnatal and adult stages. PKD2L1 expression could not be detected before E14.5 in the spinal cord at the cervical, thoracic and lumbar levels. At E14.5, few PKD2L1+ cells could be identified in the cervical spinal cord (**Figure 1A**). Starting at E16.5, PKD2L1+ cells localized around the central canal exhibited the typical morphology of spinal CSF-cNs (**Figures 1B–E**, arrows). At E18.5, the soma of PKD2L1+ cells was usually located under the layer of ependymal cells (91 out of 102, while 11 out of 102 were located within the ependyma, **Table 1**). PKD2L1+ cells sent an apical bulbous extension toward the central canal ending in the lumen with a bud enriched in PKD2L1 (**Figures 1B–E**, arrows). In the adult, the soma of PKD2L1+ CSF-cNs was located in lamina X and always under the ependyma (95 out of 95 PKD2L1+ cells) (**Figure 1E**, **Table 1**). We also observed PKD2L1+ cells with similar morphology localized away from the central canal, though these cells lacked a visible apical extension to the central canal (**Figures 1A–C**, arrowheads). Our data identify PKD2L1 as a general marker of CSF-cNs.

**FIGURE 1 | PKD2L1 expression in CSF-cNs is present from embryonic stage E14.5 to adulthood in mouse. (A–E)** Immunostaining for PKD2L1 at E14.5 **(A)**, E16.5 **(B)**, E18.5 **(C)**, P1 **(D)**, and in the adult **(E)** performed on coronal sections of the mouse cervical spinal cord. At E14.5, few PKD2L1+ cells are detected (arrowhead). From E16.5 **(B)** to adult **(E)**, multiple PKD2L1+ CSF-cNs

#### **Table 1 | Distribution of PKD2L1+ cells based on the location of their soma relative to the ependyma in the mouse spinal cord.**

*PKD2L1*+ *cells located within the ependyma represent only a small number (<11%) of the total population of PKD2L1*+ *cells counted at embryonic and newborn stages. In the adult mouse spinal cord, the somata of all PKD2L1*+ *cells are located subependymally.*

#### **PKD2L1 IS PRESENT IN GABAergic CSF-cNs IN THE MOUSE SPINAL CORD**

As mentioned before, cells contacting the CSF had been described as GABAergic in many species, but the neurotransmitter phenotype has not been thoroughly established yet in mouse. We tested whether CSF-cNs were GABAergic in the embryonic and adult mouse spinal cord (**Figure 2**). We took advantage of the wellcharacterized GAD67-GFP knock-in mouse where the enhanced GFP was targeted to the locus encoding GAD67 using homologous recombination (Tamamaki et al., 2003). In these animals, we performed IHC for GFP to amplify the endogenous GFP signals (**Figure 2**). In GAD67-GFP knock-in embryos, double IHC for GFP and PKD2L1 on coronal sections of E16.5 mice spinal cords (**Figure 2A**) revealed that most of PKD2L1+ cells (arrows) surround the central canal where they project an apical bulbous dendritic extension contacting its lumen. Note that PKD2L1+ cells exhibiting a similar morphology but not distinctly contacting the central canal (arrowheads) can also be observed **(A–C)**. Dorsal is up. White dashed line delineates the central canal. DAPI staining appears in blue. Scale bars = 20μm.

were GFP<sup>+</sup> (arrows) indicating their GABAergic nature (*n* = 2 embryos, 107 out of 162 PKD2L1+ cells). In one adult, we observed the same findings and found 47 out of 73 cells labeled for both PKD2L1 and GFP (**Figure 2B**, arrows). Note that both in the embryos and in the adult, we observed PKD2L1+cells that were not distinctly GFP+ (**Figures 2A,B**, double arrowhead). Notably, we never observed GFP+ CSF-cNs that were not PKD2L1+. Altogether we show evidence for PKD2L1+ CSF-cNs that are GABAergic in the mouse spinal cord.

#### **PKD2L1+ GABAergic CSF-cNs SURROUND THE CENTRAL CANAL IN THE ADULT MACAQUE SPINAL CORD**

We next asked whether PKD2L1 labels CSF-cNs in the spinal cord of primates (**Figure 3**). In transverse sections from cervical, thoracic and lumbar spinal cord, we observed the distribution of PKD2L1 immunofluorescence around the central canal in lamina X (**Figures 3A–C**). PKD2L1+ cells were localized around the central canal and exhibited the typical morphology of spinal CSFcNs with an apical extension toward the central canal ending with a bud in the lumen. PKD2L1 was enriched in the terminal bud of cerebrospinal fluid-contacting cells (**Figures 3A–C**) and faintly expressed (**Figures 3B,C**) or absent (**Figure 3G**) in the cell soma and in the rest of the apical extensions. PKD2L1+CSF-cNs closely surrounded the central canal; they had a round nucleus and were located under (**Figures 3B–E**) or within the ependyma (**Figures 3F,G**). In order to test whether PKD2L1+ CSF-cNs were GABAergic, we used IHC for the enzymes GAD65/67 and the VGAT transporter. Positive immunostaining for GAD65/67 and VGAT was found in cells surrounding the central canal at the level of their apical bulbous extension, soma and putative axon

**FIGURE 2 | Some PKD2L1+ cells surrounding the central canal of cervical spinal cord of adult mouse are immunoreactive for GFP in the knock-in GAD67-GFP mouse. (A)** 10μm-thick coronal sections of GAD67-GFP E16.5 mouse spinal cord immunostained for GFP **(A1,A3,A4)** and PKD2L1 **(A2–A4)**. Most PKD2L1+ cells are GFP+ (arrows) but not all

(double arrowheads). **(B)** 30μm-thick coronal sections of a GAD67-GFP adult mouse spinal cord immunostained for GFP **(B1,B3,B4)** and PKD2L1 **(B2–B4)**. While many PKD2L1+cells are distinctly GFP+ (arrows), some cells are not (double arrowheads). Dorsal is up. White dashed line delineates the central canal. DAPI staining appears in blue. Scale bars = 25μm **(A)**, 15μm **(B)**.

(**Figures 3D–F**, arrowheads, arrows, and asterisks, respectively) but buds were labeled by GABAergic markers less frequently than they were by PKD2L1. Double immunofluorescence for PKD2L1 and VGAT showed evidence for CSF-cNs double-labeled at the level of the soma and the intraluminal bud (**Figure 3G**). Nonetheless VGAT could not be detected in all PKD2L1+ buds (**Figure 3H**, double arrowheads). Cells double-labeled for PKD2L1 and VGAT were observed at the cervical, thoracic and lumbar level in the spinal cord. Taken together, our data show for the first time the presence of immunoreactive CSF-cNs containing PKD2L1 and GABAergic markers (GAD65/67 or VGAT) in the macaque lamina X. These findings demonstrate that a population of GABAergic CSF-cNs expressing PKD2L1 is present as well in the spinal cord of adult primates. Our results also support the idea that PKD2L1 is a specific marker of CSF-cNs in primates and labels more of these cells than GABAergic markers do in our experimental conditions.

#### **THE EXPRESSION OF** *pkd2l1* **mRNA IN ZEBRAFISH EMBRYO ENABLES AN EXTENSIVE QUANTIFICATION AND LINEAGE ANALYSIS OF CSF-cNs**

The developmental origin of CSF-cNs has not been established yet in mammals. To tackle the question of the developmental origin of *pkd2l1*+ cells, we took advantage of the zebrafish embryo for its transparency and the rapidity of development of the spinal cord.

#### *pkd2l1 is expressed in CSF-cNs in zebrafish*

To assess whether *pkd2l1* was expressed in CSF-cNs in zebrafish, we performed ISH using the antisense probe for *pkd2l1* (**Figure 4**). We observed the expression of *pkd2l1* in CSF-cNs along the central canal in sagittal sections of the adult spinal cord (**Figure 4A**). *pkd2l1*+ cells contacting the CSF were located ventrally and dorsally to the central canal (**Figures 4A1,A2**). In the zebrafish embryo, we investigated the 10-, 14-, 18-, 20-, and 30 somite stages (*n >* 6 embryos per condition). A weak signal in the rostral spinal cord (black arrow) could be detected at the 18-somite stage (**Figure 4B**). Gradually, the staining appeared rostro-caudally within the spinal cord (**Figures 4C,D**). Cells with dense labeling were located in two parallel rows along the ventral margin of the spinal cord (**Figures 4E,F**). Dorsal views of the spinal cord showed that these cells were located on either side of the midline (**Figure 4G**). At the larval stages, we observed that *pkd2l1*+ CSF-cNs covered the entire length of the spinal cord and the organization in two parallel rows was not distinct anymore (data *not shown*). In zebrafish, CSF-cNs (named KAs) have

**FIGURE 3 | PKD2L1+ CSF-cNs can be found as GABAergic neurons surrounding the central canal in the spinal cord of the adult macaque. (A–C)** PKD2L1 immunostaining labels intraluminal buds **(A–C)** and rarely subependymal or ependymal cell bodies **(B,C). (D,E)** GAD65/67

immunoreactive cells are numerous in the vicinity of the central canal: proximal cells show a clear apical extension contacting the lumen while cells located away do not. **(F)** CSF-cNs surrounding the central canal are *(Continued)*

#### **FIGURE 3 | Continued**

immunoreactive for the vesicular GABA transporter VGAT. **(G,H)** Double immunostaining for VGAT and PKD2L1 shows that some but not all PKD2L1+ CSF-cNs are clearly GABAergic. The intraluminal buds can be found double-labeled for PKD2L1 and VGAT (arrowheads) but not systematically (double arrowheads), while apical extensions, cell body and putative axon are

**FIGURE 4 |** *pkd2l1* **is expressed from the embryonic 18-somite stage to adulthood in the zebrafish spinal cord. (A)** Adult; **(B–G)** embryonic stages: 18-somite **(B)**, 20-somite **(C)** and 30-somite stages **(D–G)**. **(A)** In sagittal sections from the adult spinal cord, *pkd2l1*+ CSF-contacting cells are located ventral and dorsal to the central canal. White dash line: central canal, dark gray line: ventral limit of the spinal cord. *pkd2l1* expression appears in the rostral spinal cord (**B,C**, arrows) and is distributed in two rows of cells along the rostro-caudal axis **(E,F)** and on each side of the midline **(G)**. **(E)** is a close-up of the black box from **(D)**. Rostral is to the left for all panels. Lateral views with dorsal up for **(A–F)** and dorsal view for **(G)**. DAPI staining appears in blue. Scale bars = 40μm **(A)**, 100μm for **(B–D)**, 50μm for **(E–G)**.

usually only labeled by VGAT. All panels are coronal sections of the macaque adult spinal cord taken at the lumbar level for **(A,G)**, at the cervical level for **(B–F)** and at the thoracic level for **(H)**. In all panels, arrows point to the somata, asterisks to the putative axons and arrowheads to the intraluminal buds of CSF-cNs. White dashed line delineates the central canal. DAPI staining appears in blue.  KAs were therefore divided into two subpopulations on this basis, dorsal KA' and ventral KA" (Park et al., 2004). To identify the nature of *pkd2l1* expressing cells, we performed FISH in 30-somite stage zebrafish embryos (i.e., Prim-5 stage or 24 h post fertilization, **Figure 5**). This staining confirmed that expression of *pkd2l1* mRNA was localized in the ventral spinal cord and mainly distributed in two rows of cells along the rostro-caudal axis (**Figures 5A–C**). At higher magnification, we observed that these cells were lining the central canal either ventrally (arrowhead) or dorsally (arrow) (**Figures 5B–D**). Transverse sections confirmed that the somata of these cells directly surround the central canal (**Figure 5D**). While most *pkd2l1*+ cells lined the central canal, we observed faint *pkd2l1* expressing cells away from the central canal in the dorsal spinal cord (**Figures 5C,D**, double arrowhead). We quantified the number of cells for each subpopulation based on lateral views originating from 13 embryos at the 30-somite stage (**Figure 5E**). *pkd2l1*+cells localized mainly in the rostral two thirds of the spinal cord in 30-somite stage zebrafish embryos with 3–4 cells per somite for both ventral KA" and dorsal KA' between somites 5 and 15. At this stage, cell density decreased from somite 16 to 21 after which expression ceased (**Figure 5E**). We quantified the total number of *pkd2l1*+ cells for each subpopulation per embryo (**Figure 5F**). We found that *pkd2l1*+ cells from the ventral row reached 45.4 ± 2.4 cells (*n* = 13 embryos), similar to counts performed previously on KA" at the same stage (Huang et al., 2012). These observations on cell body location and number suggest that the ventral *pkd2l1*+ cells are KA" and dorsal ones are KA'.

#### *All pkd2l1***+** *CSF-cNs are GABAergic in the zebrafish spinal cord*

To demonstrate that zebrafish *pkd2l1*+ CSF-cNs are GABAergic, we investigated the expression of GABA and of GAD65/67 (**Figure 6**). An immunostaining for GABA at the 30-somite stage revealed multiple GABAergic cell types (Bernhardt et al., 1992; Higashijima et al., 2004a) including ventral and dorsal cells surrounding the central canal (**Figure 6A**, arrowhead and arrow, respectively). A short ascending axon originating from the soma of these cells can be observed (**Figure 6A**, asterisks). Based on morphology and location (see Materials and Methods), we estimated the number of GABAergic ventral (34.6 ± 2.3) and dorsal (49.9 ± 1.8) CSF-cNs per embryo (*n* = 10 embryos). We performed *pkd2l1* FISH followed by IHC for GAD65/67 to determine whether *pkd2l1*+ CSF-cNs are also GABAergic. All *pkd2l1*<sup>+</sup> cells (*n* = 160) along the central canal were GAD65/67 immunoreactive (**Figure 6B**) (92 ventral *pkd2l1*+ cells, 68 dorsal *pkd2l1*+ cells). Based on their location, density and GABAergic phenotype, these *pkd2l1*+ CSF-cNs correspond to KA" and KA'. To test whether zebrafish CSF-cNs persist as GABAergic neurons until adulthood, we performed *pkd2l1* and *gad67* double FISH on whole adult spinal cord indicating that *pkd2l1*+ CSF-cNs express *gad67* throughout development (**Figure 6C**). Contrary to what we observed in mouse and macaque, these results demonstrate that in zebrafish all *pkd2l1*+ CSF-cNs are clearly GABAergic.

#### *pkd2l1***+***CSF-cNs originate from the p3 and pMN progenitor domains in zebrafish*

To confirm the developmental origins of *pkd2l1*+ cells, we took advantage of the zebrafish model in which *pkd2l1* is expressed in the 30-somite stage embryo when progenitor domains are well-defined. Previous studies have shown that dorsal KA' cells are derived from the pMN domain and express the transcription factor *olig2* (Park et al., 2004). Ventral KA" cells originate from the p3 progenitor domain (Schafer et al., 2007) and express the transcription factor *nkx2.2a* (Yang et al., 2010; Huang et al., 2012). We therefore performed *pkd2l1* FISH combined with an anti-GFP immunostaining in the *Tg(nkx2.2a:mEGFP)* and the *Tg(olig2:EGFP)* transgenic lines (Shin et al., 2003; Ng et al., 2005) (**Figures 7**, **8**). In the *Tg(nkx2.2a:mEGFP)* transgenic embryos, the p3 domain was labeled by GFP staining as expected (**Figure 7A**). Ventral *pkd2l1*+cells contacting the central canal were labeled for GFP while dorsal cells were not (**Figure 7B**). In the *Tg(olig2:EGFP)* transgenic embryos (**Figure 8**), the progenitor domain pMN was labeled by GFP (**Figure 8A**). In these animals, only dorsal *pkd2l1*+ cells contacting the central canal were GFP+ (**Figure 8B**). Transverse sections allowed to estimate the number of *pkd2l1*+ cells contacting the central canal that were positive for GFP in the *Tg(nkx2.2a:mEGFP)* and in the *Tg(olig2:EGFP)* transgenic embryos (**Table 2**). While only ventral *pkd2l1*+cells expressed GFP in the *Tg(nkx2.2a:mEGFP)* line (38 out of 38 cells), dorsal cells solely expressed GFP in the *Tg(olig2:EGFP)* line (93 out of 93 cells) (**Table 2**). Our data indicate that *pkd2l1*+cells lining the central canal in zebrafish are KAs originating from p3 (for KA") and from pMN (for KA') depending on their dorso-ventral location relative to the central canal.

#### **DISCUSSION**

#### **PKD2L1 APPEARS AS A SPECIFIC MARKER OF SPINAL CSF-cNs IN BONY VERTEBRATE SPECIES**

PKD2L1+ neurons projecting their apical bulbous dendritic extension into the central canal of the spinal cord had been described only in mouse at the P1–P4 stages (Huang et al., 2006) and in the adult (Orts-Del'Immagine et al., 2012, 2014). We hypothesized that the channel PKD2L1 could be a specific marker of spinal CSF-cNs shared among vertebrates. We demonstrate here that the TRP channel PKD2L1 is a marker of CSF-cNs in three vertebrate species: *Mus musculus*, *Macaca fascicularis*, and *Danio rerio*. For the first time, we show the expression of this channel in primate and zebrafish CSF-cNs. In the adult macaque spinal cord, CSF-cNs express PKD2L1 at the cervical, thoracic and lumbar levels. In the zebrafish embryo, *pkd2l1*+ CSF-cNs originally differentiate in two rows along rostro-caudal gradient.

Taken together, our results demonstrate the shared expression of PKD2L1 in spinal CSF-cNs across bony vertebrate species. GABA has been previously reported to label some CSF-cNs (Barber et al., 1982; Dale et al., 1987a,b; Brodin et al., 1990; Christenson et al., 1991a,b; Bernhardt et al., 1992; Martin et al., 1998; Binor and Heathcote, 2001; Stoeckel et al., 2003; Robertson et al., 2007; Rodicio et al., 2008; Villar-Cervino et al., 2008; Reali et al., 2011). In mouse and macaque, PKD2L1 appeared always highly enriched in the intraluminal buds of CSF-cNs. On the contrary, GABAergic markers (GAD65/67, VGAT) rarely labeled in these species the soma or the apical bulbous extension of CSFcNs. Notably we never observed CSF-cNs labeled by GABAergic markers and not by PKD2L1. In mammals, PKD2L1 seems therefore to label more CSF-contacting cells than GABAergic markers do. Nonetheless, with the approach we developed here, we cannot assess the existence of CSF-cNs that would not express PKD2L1.

#### **EXPRESSION OF PKD2L1 IN CSF-cNs OF THE EMBRYONIC SPINAL CORD**

In mouse and zebrafish, we detected the expression of the channel at early embryonic stages of development (E14.5 stage for the mouse and 18-somite stage for the zebrafish). In mouse, PKD2L1 expression could be found in few spinal cells contacting the central canal at E14.5 but became more evenly expressed from E16.5 to adulthood. This observation is consistent with a recent report showing that CSF-cNs emerged at E14 in the rat spinal cord (Kutna et al., 2013). It suggests that PKD2L1 expression may start soon after the cell differentiates into a CSF-cN.

We demonstrate here that within the spinal cord of the zebrafish embryo *pkd2l1* mRNA is enriched in CSF-cNs that are mainly arranged in two rows, one ventral and one dorsal to the central canal. Previous studies have shown that KA cells can be subdivided into two populations of CSF-cNs: the ventral KA" and the dorsal KA' (Park et al., 2004; Schafer et al., 2007; Shin et al., 2007; Yeo and Chitnis, 2007; Yang et al., 2010; England et al., 2011; Huang et al., 2012). We showed that *pkd2l1* can be detected in both KA" and KA'.

In the three species studied here, we observed PKD2L1+ perikarya away from the central canal and for which we could not observe an apical bulbous extension reaching the lumen. In the zebrafish embryonic spinal cord, *pkd2l1* expression was weak in cells that were localized dorsally and away from the central canal. These cells were less than 8% (7.6% ± 2.3) of the *pkd2l1*<sup>+</sup> cells in our FISH experiments (**Figure 5**). Some of them expressed GFP in the *Tg(olig2:EGFP)* line (**Table 2**) suggesting that they could be either CSF-cNs originating from pMN, motoneurons or ventral longitudinal descending neurons (VeLDs) (Bernhardt et al., 1990; Park et al., 2004; Warp et al., 2012). A more extensive characterization would be necessary to identify the nature of these marginal dorsal cells. In cross sections of mouse, macaque and zebrafish spinal cords, those distant cells did not distinctly contact the central canal. However, as shown in rats (Lu et al., 2008), they could extend a long apical bulbous extension reaching the central canal that would be difficult to capture in thin sections.

**FIGURE 5 |** *pkd2l1* **is expressed in cells contacting the CSF and surrounding the central canal in the spinal cord of the zebrafish embryo. (A–D)** FISH of *pkd2l1* mRNA at the 30-somite stage. Lateral views for **(A–C)** and transverse section for **(D)**. Two subpopulations of *pkd2l1* bright expressing cells surround the central canal ventrally (arrowhead) and dorsally (arrow). Note the existence of dorsal *pkd2l1* weak expressing cells away from the central canal (double arrowhead). The white dashed line, dark gray solid line and the solid white lines

#### **ON THE DEVELOPMENTAL ORIGINS OF** *pkd2l1* **EXPRESSING CSF-cNs**

At the embryonic stage, the neural tube is subdivided into molecularly defined neural progenitor domains generating distinct neuronal subtypes in vertebrates (Ericson et al., 1997; Briscoe et al., 1999; Jessell, 2000; Novitch et al., 2001; Goulding, 2009). In zebrafish, CSF-cNs referred to as KA cells derive from the two most ventral domains of the spinal cord; the ventral to the central canal p3 domain labeled by *nkx2.2a* and the more dorsal pMN marked by *olig2* (Park et al., 2004; Schafer et al., 2007; Shin et al., 2007; Yeo and Chitnis, 2007; Yang et al., 2010; England et al., 2011; Huang et al., 2012). By analyzing transverse sections of stable transgenic lines where GFP reports the expression of these transcription factors, we found that *pkd2l1*+ CSF-cNs were derived from the *nkx2.2a* and from the *olig2* expression domains. This observation confirms the double developmental origin of CSF-cNs in zebrafish.

In the embryonic mouse spinal cord, the progenitor domains p3 and pMN form well-defined bands labeled by NKX2.2 and OLIG2, respectively, between E9.5 and E12.5 (Briscoe et al., 1999; Jessell, 2000; Novitch et al., 2001). At these stages, PKD2L1 is not yet expressed (*data not shown*). Therefore, by IHC for these transcription factors and PKD2L1, we could not test whether PKD2L1+CSF-cNs originate as well from p3 and pMN (*data not shown*). To reveal the developmental origin of these cells in mouse, lineage tracing based on the use of inducible transgenic lines for p3 and pMN markers will be necessary. Previous results relying on a tamoxifen-inducible Cre-recombinase inserted into indicate respectively the central canal, the ventral limit of the spinal cord, and the somite boundaries. DAPI staining appears in blue. Scale bars = 50μm for **(A)**, 30μm for **(B,C)**, and 20μm for **(D)**. **(E)** Mean number of *pkd2l1*+ cells per somite along the rostro-caudal axis at the 30-somite stage. **(F)** Total cell counts per embryo (*n* = 13 embryos). We counted 45.4 ± 2.4 ventral *pkd2l1*<sup>+</sup> cells, 47.1 ± 3.4 dorsal *pkd2l1*<sup>+</sup> cells and 7.6 ± 2.3 dorsal away from the central canal *pkd2l1*<sup>+</sup> cells per embryo. **(E,F)** Mean values are given ± s.e.m.

the Olig2 locus indicate that a subpopulation of cells originating from pMN between E9.5 and E14.5 are located at the ependymal border circling the central canal (Srinivas et al., 2001; Masahira et al., 2006). This observation suggests that some CSF-cNs could originate from pMN in mouse, although a thorough investigation would be necessary to address this question in this model organism.

 Here we used different GABAergic markers, GABA itself, the synthesis enzyme GAD or the GABA transporter VGAT, to test whether PKD2L1+ CSF-cNs were GABAergic. In zebrafish, we demonstrate that all *pkd2l1*+ cells are GABAergic in spinal CSF-cNs. In mouse and macaque, we could not demonstrate a complete co-localization of PKD2L1 with GABAergic markers (GAD67-GFP or VGAT) in CSF-cNs. In macaque, a minority of PKD2L1+ intraluminal buds was clearly double-labeled with VGAT. In mouse, the observation of GABAergic PKD2L1+ CSF-cNs was confirmed in the GAD67- GFP transgenic line by the co-expression of PKD2L1 and GFP in only some cells. The lack of systematic colocalization between PKD2L1 and GABAergic markers in CSF-cNs could be due to

the expression of other isoforms of GAD (such as GAD65) or to other neurotransmitters expressed in PKD2L1+ CSF-cNs. Although we could not demonstrate that GABA is expressed in all CSF-cNs in mammals, our results show evidence for GABAergic PKD2L1+ CSF-cNs in the spinal cord of the three species studied here.

Previous studies characterized a diversity of markers potentially expressed in CSF-cNs in multiple species. Peptides such as the vasoactive intestinal polypeptide (VIP), somatostatin or urotensin II-related peptide (URP2) have been found in CSF-cNs (Buchanan et al., 1987; Yulis and Lederis, 1988a,b; Christenson et al., 1991a; Lamotte and Shapiro, 1991; Lopez et al., 2007; Wyart et al., 2009; Parmentier et al., 2011; Jalalvand et al., 2014). Ventral CSF-cNs were found dopaminergic in tetrapods such as birds (Acerbo et al., 2003) and amphibians (Gonzalez and Smeets, 1991, 1993; Gonzalez et al., 1993), as well as in some teleosts such as the eel and the trout (Roberts et al., 1995), in dogfish (Sueiro et al., 2004) and lampreys (Schotland et al., 1996; Rodicio et al., 2008). It would be interesting to determine in these species whether the dopaminergic and non-dopaminergic CSF-cNs derive as well from two different progenitor domains. However, the dopaminergic phenotype does not seem to be conserved in all vertebrate species as shown in some teleosts and in mammals (Nagatsu et al., 1988; McLean and Fetcho, 2004).

Other markers such as the subunit P2X2 of the purinergic receptor have been found in adult CSF-cNs together with early neuronal markers such as the polysialylated neural cell adhesion molecule (PSA-NCAM) in rats (Stoeckel et al., 2003) and HuC in turtles (Reali et al., 2011) and dogfish (Sueiro et al., 2004).

All markers listed above are not highly specific for CSF-cNs. However, in the three species studies here, PKD2L1 appears as a specific marker of CSF-cNs that is highly expressed and broadly targets CSF-cNs within the spinal cord. Despite existing variations in expression of peptides, neuromodulators and receptors, our results in zebrafish, mouse and macaque suggest the existence of a conserved system of spinal CSF-cNs defined by their morphology, their location—ependymally or subependymally—and their enriched expression of PKD2L1.

**FIGURE 7 | Ventral** *pkd2l1***+ cells express** *nkx2.2a* **and derive from the p3 progenitor domain in the zebrafish embryo. (A,B)** *pkd2l1* FISH in *Tg(nkx2.2a: mEGFP)* embryos immunostained for GFP at the 30-somite stage. IHC for GFP reveals the p3 domain. **(A)** Lateral view showing that the most ventral *pkd2l1*+ cells are GFP+. **(B)** A typical transverse section shows

a ventral *pkd2l1*+ cell contacting the central canal and expressing GFP (arrowhead) while a dorsal *pkd2l1*+ cell contacting the central canal does not express GFP (arrow) in the *Tg(nkx2.2a: mEGFP)* transgenic embryo. White dashed line delineates the central canal, dark line the ventral limit of the spinal cord. DAPI staining appears in blue. Scale bars = 20μm.

**FIGURE 8 | Dorsal** *pkd2l1***+ CSF-cNs express** *olig2* **and derive from the pMN progenitor domain in the embryonic spinal cord of zebrafish. (A,B)** *pkd2l1* FISH in *Tg(olig2:EGFP)* embryos immunostained for GFP at the 30-somite stage. IHC for GFP reveals the pMN domain. **(A)** Lateral view showing dorsal *pkd2l1*+ cells express GFP. **(B)** Transverse sections showing

dorsal *pkd2l1*+ cells contacting the central canal express GFP (arrow) while a ventral *pkd2l1*+ cell contacting the central canal (arrowhead) does not express GFP in the *Tg(olig2:EGFP)* transgenic line. White dashed line delineates the central canal, dark line the ventral limit of the spinal cord. DAPI staining appears in blue. Scale bars = 20μm.

**Table 2 | Distribution of ventral and dorsal** *pkd2l1***+ cells expressing GFP in** *Tg(nkx2.2a:mEGFP)* **and** *Tg(olig2:EGFP)* **zebrafish transgenic lines at the 30-somite stage.**

*All ventral pkd2l1*+ *cells are GFP*+ *in Tg(nkx2.2a:mEGFP) but none of the dorsal ones. Reciprocally all the dorsal pkd2l1*+ *cells are GFP* + *in the Tg(olig2:EGFP) but none of the ventral ones.*

 The peculiar location of CSF-cNs in contact with the CSF strongly suggests a role for PKD2L1 as a sensor of CSF composition, pH and/or osmolarity (Huang et al., 2006; Orts-Del'Immagine et al., 2012) since the channel is activated upon acidification (Ishimaru et al., 2006; Inada et al., 2008; Orts-Del'Immagine et al., 2012), alkalinization (Shimizu et al., 2011; Orts-Del'Immagine et al., 2012) or hypoosmotic variations (Shimizu et al., 2009; Orts-Del'Immagine et al., 2012).

The role(s) of CSF-cNs in the vertebrate spinal cord is (are) poorly understood. They could be proprioceptors sensitive to the CSF composition (Huang et al., 2006; Orts-Del'Immagine et al., 2012) and modulating locomotion (Wyart et al., 2009), or enabling the differentiation of progenitors in the ependymal neurogenic niche via GABA release (Reali et al., 2011). The investigation of PKD2L1 functions in CSF-cNs across multiple species should reveal whether its physiological role in the spinal cord is conserved in vertebrates.

#### **AUTHOR CONTRIBUTIONS**

Lydia Djenoune and Claire Wyart conceived, designed, and supervised all experiments. Lydia Djenoune performed the experiments on zebrafish embryos helped by Céline Burcklé at the early stages of the project. Hanen Khabou performed all experiments on the macaque spinal cord. Fanny Joubert and Laurence Bodineau performed experiments on the mouse spinal cord. Feng B. Quan performed FISH experiments on the zebrafish adult spinal cord. Sophie Nunes Figueiredo performed GABA immunohistochemistry on zebrafish embryos. Filippo Del Bene designed the *pkd2l1 in situ* probe. Lydia Djenoune analyzed the data under supervision of Claire Wyart and Hervé Tostivint. Lydia Djenoune and Claire Wyart wrote the manuscript. All authors discussed the results and implications and commented on the manuscript.

#### **ACKNOWLEDGMENTS**

We thank Prof. Bruce Appel, University of Colorado in Denver, USA for sharing the *Tg(nkx2.2a:mEGFP)* and *Tg(olig2:EGFP)* fish transgenic lines and Prof. Yuchio Yanagawa, Gunma University Graduate School of Medicine in Maebashi, Japan for sharing the GAD67-GFP knock-in mice used in this study. We thank Dr. Uwe Strähle, Karlsruhe Institute of Technology, Germany for sharing the *gad67* plasmid. We thank Jean Simonnet, Dr. Desdemona Fricker, and Dr. Richard Miles for help with the GAD67-GFP knock-in mouse. We thank Dr. Vanessa Ribes for helpful advices regarding experiments on the embryonic mouse spinal cord. We thank Dr. Carlos Parras, Sowmya Sekizar, Dr. Mariana Graciarena, Melissa Fauveau, and Dr. Brahim Nait-Oumesmar who gave valuable insights for IHC in the mouse embryonic spinal cord. We thank Dr. Pierre Pouget and Prof. Marie Vidailhet for providing access to the tissue of adult macaque spinal cord and Virgile Brochard and Dominique Tandé for precious advices. We thank Prof. Herwig Baier, Max Planck Institute of Neurobiology in Martinsried, Germany, the team of the "Plateforme d'Imagerie Cellulaire Pitié Salpêtrière" and Prof. Thomas Similowski for their support. We thank Urs Lucas Böhm, Kevin Fidelin, Jenna Sternberg, Dr. Pierre-Luc Bardet, Dr. Andrew E. Prendergast, Dr. Jean Paul Rio, and Dr. Kristen Severi for critical reading of the manuscript. This work received financial support from the Institut du Cerveau et de la Moelle épinière (ICM with the French program "Investissements d'avenir" ANR-10-IAIHU-06), the network Ecole des Neurosciences de Paris (ENP), the Fondation Bettencourt Schueller (FBS), Mr Pierre Belle, the City of Paris Emergence program, the Atip/Avenir junior program from Institut National de la Santé et de la Recherche Médicale and Centre National de la Recherche Scientifique, the Fyssen foundation, the International Reintegration Grant from Marie Curie Actions Framework Program 6, and the European Research Council (ERC) starter grant "OptoLoco."

#### **REFERENCES**

**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.

*Received: 05 January 2014; accepted: 10 April 2014; published online: 06 May 2014. Citation: Djenoune L, Khabou H, Joubert F, Quan FB, Nunes Figueiredo S, Bodineau L, Del Bene F, Burcklé C, Tostivint H and Wyart C (2014) Investigation of spinal cerebrospinal fluid-contacting neurons expressing PKD2L1: evidence for a conserved system from fish to primates. Front. Neuroanat. 8:26. doi: 10.3389/fnana.2014.00026 This article was submitted to the journal Frontiers in Neuroanatomy.*

*Copyright © 2014 Djenoune, Khabou, Joubert, Quan, Nunes Figueiredo, Bodineau, Del Bene, Burcklé, Tostivint and Wyart. 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) or licensor 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.*

#### *Filipe Pinto-Teixeira1,2†, Mariana Muzzopappa1, Jim Swoger 1,2, Alessandro Mineo1,2, James Sharpe1,2,3 and Hernán López-Schier 1,4\**

*<sup>1</sup> Centre for Genomic Regulation (CRG), Barcelona, Spain*

*<sup>2</sup> Universitat Pompeu Fabra (UPF), Barcelona, Spain*

*<sup>3</sup> Institució Catalana de Recerca i Estudis Avançats (ICREA), Barcelona, Spain*

*<sup>4</sup> Unit Sensory Biology and Organogenesis, Helmholtz Zentrum München, Munich, Germany*

#### *Edited by:*

*Laurent Gautron, UT Southwestern Medical Center, USA*

#### *Reviewed by:*

*Kazuhide Asakawa, National Institute of Genetics, Japan Mireille Montcouquiol, INSERM, France*

#### *\*Correspondence:*

*Hernán López-Schier, Unit Sensory Biology and Organogenesis, Helmholtz Zentrum München, Ingolstaedter Landstrasse 1, 85764 Munich, Germany e-mail: hernan.lopez-schier@ helmholtz-muenchen.de*

#### *†Present address:*

*Filipe Pinto-Teixeira, Department of Biology, New York University, New York, USA*

Direct videomicroscopic visualization of organ formation and regeneration *in toto* is a powerful strategy to study cellular processes that often cannot be replicated *in vitro*. Intravital imaging aims at quantifying changes in tissue architecture or subcellular organization over time during organ development, regeneration or degeneration. A general feature of this approach is its reliance on the optical isolation of defined cell types in the whole animals by transgenic expression of fluorescent markers. Here we describe a simple and robust method to analyze sensory hair-cell development and regeneration in the zebrafish lateral line by high-resolution intravital imaging using laser-scanning confocal microscopy (LSCM) and selective plane illumination microscopy (SPIM). The main advantage of studying hair-cell regeneration in the lateral line is that it occurs throughout the life of the animal, which allows its study in the most natural context. We detail protocols to achieve continuous videomicroscopy for up to 68 hours, enabling direct observation of cellular behavior, which can provide a sensitive assay for the quantitative classification of cellular phenotypes and cell-lineage reconstruction. Modifications to this protocol should facilitate pharmacogenetic assays to identify or validate otoprotective or reparative drugs for future clinical strategies aimed at preserving aural function in humans.

**Keywords: hair cells, auditory, zebrafish, intravital fluorescence microscopy, regeneration, lateral line system, development**

#### **INTRODUCTION**

Direct *in toto* visualization of organ formation and repair is a powerful strategy to study cellular processes that often cannot be replicated *in vitro* (Megason and Fraser, 2007; Pittet and Weissleder, 2011; Rompolas et al., 2012). A general feature of high-resolution intravital imaging is its reliance on the optical isolation of cells or sub-cellular structures in the whole animal by the transgenic expression of fluorescent markers (Faucherre et al., 2010; Dempsey et al., 2012). The formation of specialized organs during embryonic development involves the activation of genetic programs that determine cell fate, cell proliferation, and the assembly of tissues into functional organs. Animals spend the overwhelming majority of their lives as adults, when many organs are under acute or chronic physical or toxicological stress and can sustain damage. Most organisms have evolved two general strategies to maintain organ function throughout life. The most prevalent is protection by isolating organs into body cavities. Although effective, isolation is not optimal for sensory systems because their external components need to be exposed to the environment to fulfill their function. Therefore, organisms have evolved a second strategy to maintain sensory abilities over long periods: regeneration and repair. Paradoxically, however, mammals have lost the capacity to repair the sensory elements of the inner ear (Corwin and Oberholtzer, 1997; Kelley, 2003; Kros, 2007; Edge and Chen, 2008; Brigande and Heller, 2009; Rubel et al., 2013). If damaged by infection, drugs or stress, the ear's mechanosensory hair cells cannot be replaced, leading to irreversible loss of hearing and balance (Collado et al., 2008). The resulting sensory dysfunction develops into a major handicap that dramatically decreases the quality of life of the affected individuals. Deafness often causes acute communication difficulties, which represents a significant socioeconomic problem considering that it afflicts more than 10% of the population in industrialized countries (Magilvy, 1985). According to the British charity "Deafness Research UK," 1 in 7 people in the United Kingdom suffer from hearing impairment. Notwithstanding the biological and clinical interest of deafness and balance disorders, the mechanisms that underlie hair-cell homeostasis remain poorly understood because of the inherent difficulty to study the inner ear in animals after birth. By contrast, aquatic vertebrates such as the zebrafish have a superficial and anatomically simple "hearing" system called lateral line (**Figures 1A–C**) (Raible and Kruse, 2000; Ghysen and Dambly-Chaudiere, 2007; Behra et al., 2009). This mechanosensory system detects hydromechanical variations around the animal body and serves to command behaviors such as schooling, rheotaxia, prey capture, and obstacle and predator avoidance (Dijkgraaf, 1963; Ghysen and Dambly-Chaudiere, 2007; Bleckmann and Zelick, 2009). Fishes of lotic and lentic freshwater ecosystems have a welldeveloped superficial lateral-line system, whose functional units

Low magnification view of a zebrafish larva under transmitted light. **(B)** Hair cells in the lateral line were labeled with the fluorescent vital dye DiAsp (bright orange). It shows the superficial distribution of neuromasts along the anterior (head) and posterior (trunk and tail) lateral-line systems. **(C)** Schematic representation of the distribution of the neuromasts in the lateral line of a zebrafish larva. Orange highlights the anterior neuromasts,

green the posterior neuromasts, and blue the dorsal neuromasts. **(D,E)** High magnification confocal image of a frontal view of a posterior neuromast revealing the hair cells (blue), supporting cells (red and green). **(F)** Schematic representation of a neuromast viewed from the side, depicting every known cell, including the neurons. **(G)** High magnification side-view image of a mature neuromast labeled with the vital dye DiAsp. Scale bars are 10μm.

are called neuromasts (**Figures 1D–G**). The superficial lateral line is very accessible to experimentation and imaging. The neuromast is a volcano-shaped epithelium that projects from the animal's body surface, into the water (**Figure 1F**). A mature neuromast is 20μm in height and has a diameter of 50μm. It is formed by around 60 cells, of which one third are hair cells and the remainder divided between peripheral mantle cells and central sustentacular (supporting) cells (**Figures 1D,E**). Importantly, identically to their mammalian counterparts, hair cells in the zebrafish are susceptible to ototoxicity or mechanical damage (Harris et al., 2003; Chiu et al., 2008; Behra et al., 2009; Brignull et al., 2009). However, unlike mammals, fishes can regenerate hair cells rapidly and orderly after damage (Williams and Holder, 2000; Harris et al., 2003; López-Schier and Hudspeth, 2006; Behra et al., 2009). Our group was the first to unambiguously identify resident haircell progenitors, and to study how their dynamic relationship with supporting cells maintains a constant, functional organ throughout the animal's life (López-Schier and Hudspeth, 2006). Our results have revealed that hair cells in neuromast are always born sequentially and in pairs along a single axis (**Figure 2**) (López-Schier and Hudspeth, 2006). These observations allowed us to predict the compartmentalization of the neuromast epithelium. We subsequently confirmed this prediction using the SqET4 transgenic zebrafish line that identified resident hair-cell progenitors, and the SqET20 line that revealed a cellular territory where hair-cell progenitors generate at high frequencies (Wibowo et al., 2011). Thus, the detailed intravital *in vivo* analyses have unveiled two previously unknown cell types in neuromasts. Here we detail a simple, fast and inexpensive protocol to characterize hair-cell development and regeneration by *in toto* high-resolution live imaging. This protocol has been optimized during over 10 years of experience using the zebrafish and high-resolution imaging (López-Schier and St. Johnston, 2001; López-Schier et al., 2004; López-Schier and Hudspeth, 2006; López-Schier, 2010; Swoger et al., 2011; Wibowo et al., 2011). It relies on the stable expression of engineered transgenes coding for fluorescent proteins in specific cellular populations of the lateral line. The procedure only requires a small zebrafish colony and access to a Laser-Scanning Confocal Microscope (LSCM) for image acquisition (Wibowo et al., 2011). We also describe the handling and mounting of samples to enable imaging by Selective Plane Illumination Microscopy (SPIM) (Huisken et al., 2004; Greger et al., 2007; Huisken and Stainier, 2009; Swoger et al., 2011; Kaufmann et al., 2012). Finally, this approach requires a desktop computer equipped with free or inexpensive commercial software for qualitative or quantitative image analysis and rendering.

#### **RATIONALE**

Hair-cell development is likely regulated by mechanisms that are cell-type specific, and controlled by a complex activation of signaling pathways in both hair-cell progenitors and supporting cells (López-Schier, 2004). Notwithstanding its major scientific and clinical significance, the mechanisms that underlie the normal production of hair cells remain poorly understood. Studies aimed at characterizing the behavior of endogenous haircell progenitors should be carried out in animal model systems with accessible neuroepithelia and capable of regenerating hair cells efficiently. Pharmacological treatments that produce deafness in mammals are also lethal to hair cells in the zebrafish. Hair cells in the zebrafish lateral line begin to regenerate less than 12 h after loss, to eventually produce a complete anatomical and functional recovery of the organ in less than 72 h. Many important questions remain unanswered: What is the source of newly formed hair cells? How is the process of hair-cell regeneration controlled? How the three-dimensional architecture of the epithelium recovers? Results from experiments in several species suggest that hair cells regenerate from supporting cells by trans-differentiation. Other results indicate that supporting cells re-enter mitosis before producing new hair cells. Although supporting cell trans-differentiation may be a simple mechanism to replace damaged hair cells, supporting-cell amplification is also required because animals need to maintain the number and proportion of the different cell types over many years. Thus, cell proliferation is eventually essential for neuroepithelial homeostasis. An attractive possibility is that the neuroepithelium harbors a population of pluripotent progenitor cells dedicated to regenerate hair cells (Harris et al., 2003; Huisken et al., 2004; Rivolta et al., 2006; Oshima et al., 2009, 2010). This hypothesis has taken some impulse from a series of results suggesting the existence of stem cells in the utricle. These cells could be characterized further for potential clinical applications. Stem-cell-based therapies represent one promising avenue to repair damaged inner ears (Brigande and Heller, 2009; Chen et al., 2012; Rivolta, 2013). However, they require invasive surgical procedures and postoperative follow-up. The shortcomings of cell replacement therapy suggest that mobilizing resident progenitor cells to replace damaged tissues is a viable alternative with therapeutic potential (Mizutari et al., 2013). This protocol provides a simple and robust method to obtain information about the molecular profile, localization, and behavior of progenitor cells within a mature organ.

**FIGURE 2 | UHCPs division and planar cell inversion.** Frames from Supplementary Movie 1 at the time-points indicated A 916 min series of confocal images of a double-transgenic Tg[Cldnb:mGFP;SqET4] regenerating neuromast. Two prospective UHCPs were identified retrospectively by playing the time series backwards and labeled with a yellow and green dot. Each UHCP eventually divides into a pair of sister hair cells, labeled in Cyan/Red and Green/Orange, respectively. On minute

624 the sibling hair cells in the lower part of the neuromast begin to rotate around their contact point, eventually breaking the line of mirror symmetry. This complete inversion of the sister hair cells restores the line of mirror symmetry realigning the cells along the neuromast axis of planar polarity (double-headed arrow in the first panel). All images correspond to a single focal plane with the cells of interest in focus. Scale bar is 10μm.

#### **THE ZEBRAFISH AS AN EXPERIMENTAL MODEL SYSTEM**

The zebrafish (*Danio rerio*) is a small tropical fish whose biology has been extensively characterized. Interesting developmental and behavioral aspects of the zebrafish have made it a favorite research organism in genetics, biomedicine, and drug discovery for the past three decades. It compares favorably with other experimental animal model systems, offering the following advantages:

#### **METHODS**

#### **HANDLING OF THE ZEBRAFISH**

The success of these procedures crucially depends on the proper handling and mounting of the zebrafish embryo or larva. Sample handling needs to be practiced to ensure animal survival. We routinely monitor the animal's vital signs after imaging (heart beating and blood flow). If samples will be further processed for immunohistochemistry staining after live imaging, they should be fixed as soon as possible after imaging ceases.

#### **OBTAINING ZEBRAFISH LARVAE FOR EXPERIMENTS**

Zebrafish are maintained in E3 Embryo medium. To make an E3 medium stock solution (60×) dissolve 172 g NaCl, 7.6 g KCl, 29 g CaCl2.2H2O and 49 g MgSO47H2O in 10 L of distilled H2O (dH2O). To avoid water spoilage add methylene blue (Sigma-Aldrich; cat. No.M4159). Make a methylene blue stock solution of 4 mgml−<sup>1</sup> in water. For a final working solution, add 70μl of stock solution to 1 L of E3 Embryo medium. Fish used were maintained under standardized conditions and experiments were performed in accordance with protocols approved by the PRBB Ethical Committee of Animal Experimentation.

#### **NECESSARY EQUIPMENT FOR RAISING FISH AND COLLECTING EGGS**

Divisible acrylic tanks for zebrafish breeding and plastic strainers for egg or larva collection. Incubator set at 28*.*5◦C to maintain embryos and larvae at constant temperature. An epifluorescenceequipped stereomicroscope for initial screening of experimental animals. In addition, disposable 6 or 12 well culture plastic plates, Pasteur plastic pipettes will be needed.

#### **HAIR-CELL ABLATION IN ZEBRAFISH LARVAE**

A stock solution of Neomycin 500 mM (neomycin trisulphate salt hydrate, SIGMA-Aldrich N6386) should be maintained at 4◦C, and diluted in E3 medium without methylene blue to a working concentration of 250μM. The stock solution should be kept in a 1.5 ml Eppendorf tube sealed with Parafilm and saved in an Eppendorf box to minimize the risk of contamination. It is important to be cautious with the use of neomycin. Mechanosensitive hair cells, including those of humans, are extremely sensitive to aminoglycosides. Neomycin is quickly absorbed by the body surface, therefore, gloves and protective goggles should be used during all the steps that involve the use of neomycin.

#### **COMPONENTS FOR MOUNTING ZEBRAFISH LARVAE**

To anesthesize zebrafish, prepare a stock solution of tricaine (MS222 or MESAB: 3-amino benzoic acidethylester) at 25×. Dissolve 0.4 g Tricaine, in 97.9 mL of dH2O. Adjust the pH to ∼7 by adding 9.1 mL of 1 M Tris (pH 9). Store at 4◦C. Make one percent low–melting point agarose in E3 to mount embryos for observation. In an Erlenmeyer flask make 50 ml of 1% LMP agarose 436 (Bio Rad, Cat. 161-3112) by adding 500 mg of agarose to 50 ml 437 of E3 medium without methylene blue. Heat the solution for 2 min in a microwave at maximum power to dissolve the agarose. If small aggregates are still found agitate the solution and heat up to obtain a clear solution. The solution will be very hot. Use a temperature protective gear to handle the Erlenmeyer flask at this stage. Transfer the solution to a 50 ml Falcon tube and store in a water bath at 42◦C. Let the agarose cool down to 42◦C before using it. This agarose can be used for one week. For confocal microscopy, hair loops are appropriate to manipulate the zebrafish larvae on cover-glass-bottomed culture dishes. For SPIM imaging, 100/200μL glass capillaries (cat. no. 701910) and piston rods (cat. no. 701936) from Brand GmbH.

#### **OBSERVATION AND ANALYSIS OF HAIR-CELL DEVELOPMENT AND REGENERATION**

A fluorescence stereomicroscope equipped with an external light source for fluorescence excitation and appropriate filter set for GFP and RFP detection can be used for screening the samples to be imaged.

For confocal imaging, a microscope equipped with lasers for GFP (488 nm), RFP (543 nm) excitation and preferentially with a motorized stage to allow multi-position imaging.

For SPIM imaging, the microscope can be equipped with lasers operating at 488 nm (for excitation of GFP) and 543 nm (RFP), and the appropriate emission filters. Excitation power and wavelength are computer-controlled via an Acousto-Optic Tunable Filter (AOTF); the filter wheel for changing emission filters is also computer controlled. Typically water-dipping objective lenses are used for detection, ranging from 20 × */*0*.*5 to 63 × */*0*.*9, depending on the desired resolution and field of view. Illumination is with air objectives, e.g., a 10 ×*/*0*.*3. Detection is performed with a CCD or sCMOS camera (in this work we assume a Hamamatsu Orca-ER CCD camera). The sample is mounted on a computer-controlled mechanical stage with 3 spatial and one rotational degrees of freedom (the rotation axis is perpendicular to the illumination and detection axes). See Greger et al. (2007)for a technical description of a SPIM microscope similar to the one used in the present work (Greger et al., 2007). We routinely use the public domain ImageJ/Fiji software for image processing.

#### **IDENTIFICATION OF TRANSGENIC EXPRESSION OF FLUORESCENT PROTEINS IN THE ZEBRAFISH LATERAL LINE**

Collect the zebrafish larvae to be screened in a Petri dish with 50 ml of methylene-blue-free E3 medium with tricaine (250μM). Screen and select embryos with a stereomicroscope using the appropriate light filter and select those larvae expressing fluorescence in the lateral line neuromasts. If the larvae to be analysed are younger than 3 dpf they have to be dechorionated. We do not recommend dechorionation with pronase as this decreases the survival rate. We favour manual dechorionation. This can easily be achieved with the help of a pair of microsurgical forceps. Hold the chorion with one forceps and with the other forceps grip the chorion and tear it. It is important to minimize the exposure of the larvae to tricaine. If the larvae are not to be treated with neomycin for the next hours transfer them to fresh methylene-blue-free E3 medium without tricaine.

#### **HOW TO IDENTIFY TRANSGENIC LARVAE USED IN THIS STUDY**

#### **NEOMYCIN INDUCED HAIR CELL ABLATION**

This step takes about 3 h. In a 100 ml Erlenmeyer flask prepare 50 ml of E3 medium without methylene blue with neomycin (250μM) by adding 250μl of the neomycin stock to 50 ml of E3 medium. Carefully agitate the Erlenmeyer to mix and transfer the neomycin solution from the Erlenmeyer to a 50 ml Petri dish. Additionally prepare one Petri dish with 50 ml of E3 medium without methylene blue. Collect the larvae in a 50 ml Petri dish with E3 medium without methylene blue with tricaine (250μM). A critical step is to keep the larvae in E3 medium with methylene blue until the desired developmental stage and transfer them to methylene-blue-free E3 medium at least 2 h before the neomycin treatment, as methylene blue affects the survival of the larvae upon treatment. With the help of a plastic Pasteur pipette transfer the larvae with the minimum amount of medium to the Petri dish with the neomycin solution. Carefully disperse the larvae in the plate by gently squeeze the Pasteur pipette. To collect the larvae hold the Petri dish on the bench and move it horizontally in very small circles until the larvae gather centripetally in the center of the petri dish. Another possibility is to use the Pasteur pipette to promote flow of the medium. This can be done by placing the tip of the pipette parallel and next to the inside border of the petri dish and gently squeeze. With the help of a clean plastic Pasteur pipette transfer the larvae with the minimal amount of medium to the previously prepared petri dish with 50 ml of E3 medium without methylene blue and let the larvae recover for at least 2 h. Some larvae will eventually die due to the neomycin treatment. These larvae should be removed from the medium.

Mechanosensitive hair cells, including those from humans, are extremely sensitive to neomycin. Use gloves throughout these steps.

#### **MOUNTING OF LARVAE FOR LIVE CONFOCAL IMAGING**

In a 1.5 ml Eppendorf tube prepare 1 ml of 1% LMP agarose with tricaine by adding 60μl of the tricaine stock solution to 940μl of the agarose solution. Keep the Eppendorf at 37◦C in a bench thermomixer. Transfer the zebrafish larvae directly to a coverglass-bottomed culture dish. With a plastic Pasteur pipette release a drop of the agarose solution to cover the larvae. Carefully rotate

the larvae with a hair loop so that the desired side to image faces the bottom side of the holder. The larvae should be mounted all in the same orientation and in the center of the dish (**Figure 3**). This step is essential if the imaging needs to be done at the early stages of neuromast development. No more than 3 larvae should be mounted per culture dish to maximize survival rate. Wait for a few minutes to let the agarose solidify (∼35 min at room temperature). When solidified, the agarose becomes a translucent and hard gel, which can be tested by touching it with the hair loop. At this stage carefully fill the dish with 2.5 ml of E3 medium without methylene blue and tricaine (250μM). Cover the dish with its lid.

#### **MOUNTING OF LARVAE FOR LIVE SPIM IMAGING**

In a 1.5 ml Eppendorf tube prepare 1 ml of 1% LMP agarose with tricaine by adding 60μl of the tricaine stock solution to 940μl of the agarose solution. Keep the Eppendorf at 37◦C in a bench thermomixer. Prepare a p20 pipette with a yellow tip by cutting 0.5 cm from the end, so that the hole is big enough for the larvae to enter. Place the capillary on the surface of the liquid agarose and pull the plunger back until the capillary is filled with 1 cm of agarose. (**Figure 4**). Immediately take the anesthetized larvae in E3 medium with the p20 pipette and carefully inject it in the capillary with the minimal amount of liquid that is possible. Lay the capillary horizontal for a few minutes to let the agarose solidify (10 min at room temperature). Mount the capillary in the SPIM device and fill the sample chamber with methylene-blue-free E3 medium with tricane.

#### **IMAGING SETUP FOR CONFOCAL MICROSCOPY**

This step takes about 1 h. To minimize photobleaching, evaporation of the medium and "burning" the larvae, the lasers should be set to the lowest settings that allow a sharp visualization of the neuromasts. As a guideline, in a Leica SP5 system laser configuration window set the Argon laser power to 30%. In the laser power adjustment window located in the beam path screen set the laser power intensity between 13% and 19%. Place the dish covered with its lid in the appropriate support. Place the dish so that the larvae are horizontally aligned in the computer acquisition screen. Locate the neuromast with a low magnification objective (20×). Use the microscope oculars to screen the larvae and identify potential neuromasts of interest. Save their location in the software location panel. From this moment, it is crucial not to touch the dish. A typical experiment allows the imaging of 4 neuromasts with a 2-min acquisition time. We recommend to save the location of 8–10 neuromasts at this pre-screening stage.

#### **NEUROMAST IDENTIFICATION**

The identification of a neuromast in the *Tg[SeqEt4]* upon the neomycin treatment, can typically be done by the presence 1– 4 hair cells (**Figure 5**). These correspond to hair cells that were not mature during the treatment, and were therefore insensible to neomycin. During lateral line development, when a neuromast is deposited a UHCP is typically dividing giving rise to two hair cells. In both cases the presence of these hair cells makes the identification of the neuromasts easier, even with a 20× objective. However, if the researcher wishes to image neuromasts where no hair cells are present, the screening has to be done using a 40× objective. In this case favor the use of *Tg[Cldnb:mGFP; SqET4]* larvae where neuromast identification can be easily done by the expression of *Tg[Cldnb:mGFP]*. The identification of neuromasts without hair cells during regeneration in the *Tg[SeqEt4]* is significantly more difficult. In this case the neuromast has to be identified using bright field light. As a guideline, neuromasts are typically located at the somite boundaries, in the middle of the myoseptum where the chevron shifts orientation (**Figure 5**). If the researcher wants to image the first pair of hair cells being produced it is necessary to start the time series when a neuromast is being deposited. The researcher

should then identify the migrating primordium, which is easily identifiable as a group of ∼100 cells just under the epidermis (**Figure 5**).

Change to a higher magnification lens (40×). For long imaging sessions an Oil Immersion objective must be used to avoid evaporation. Adjust the pixel configuration to 512 × 1024. If imaging has to be done at the early stages of neuromast development it is essential that larvae are positioned with the lateral line aligned along the long axis of the field of view, because the neuromasts are still being deposited and can move significantly during the first hours of the time lapse. Proper orientation of the lateral line avoids having the neuromasts moving out of the field of view as they are deposited. Using the previously saved locations screen the samples and identify the neuromasts you want to image. Delete undesired locations. A typical experiment allows the simultaneous imaging of 4 neuromasts.

Adjust the Z-stack for the first location. Use existing hair cells or *Tg[Cldnb:mGFP]* expression to identify the Z-stack limits. Set up the Z-stack so that it contains the entire extent of the neuromast. The Z-stack should start on the tip of the hair cells and end in the bottom part of the neuromast. As default use a step size of 2μm. If using the *Tg[Cldnb:mGFP]* with the purpose of cell tracking do not use a step size bigger than 2μm. Because hair cells develop/regenerate the neuromast assumes its typical volcano shape and its volume changes as it grows in size. This, and the continuous development of the larvae, might originate movement of the region of interest out of the Z-stack boundaries during data acquisition. To avoid this add an extra 2μm to the boundary that sets the beginning of the stack and 5 μm to the boundary that sets the end of the stack on the bottom of the neuromast/hair-cells. If the system does not allow specific Zstacks for individual positions use as general stack boundaries the tip of the hair cells/neuromast that is closest to the objective and the bottom of the neuromast/hair-cell that is furthest away from the objective. If imaging a *Tg[SeqEt4]* larvae where no hair cells are present use the skin of the animal as a reference to set the beginning boundary and from there set a ∼20μm stack. Adjust the position of the neuromast to the center of the field of view and resave the position with the new settings. Repeat this procedure for each position to be imaged.

Adjust the Gain settings. To visualize UHCP's that weakly express GFP, set up the gain so that the peak GFP signal from existing hair cells is just above the saturation threshold. If imaging neuromasts without hair cells use hair cells of another neuromast in the sample as a reference. Activate the Bidirectional Scanning option to double the scanning speed, run a trial scan and judge the quality of the images. Adjust the acquisition time and length according to the needs. If the acquisition time is less than the total time between the start of consecutive time-points, it might be ideal to apply an increased number of averaging passes to improve image quality. For live imaging it is always recommended to use an average between lines to avoid any image disturbance due to movement of the tissue. Save the LIF file and then save each time-lapse as individual TIFF series with the name "STACKNAME\_STACK TIFFS." Create a logbook with the following details of each time lapse: Transgenic line used, duration of time lapse, Z-stack step size, Z-stack number of steps.

#### **IMAGING SETUP FOR SPIM**

The primary advantage for live-sample imaging that SPIM has over confocal microscopy is the reduced photo-damage of the sample. In confocal microscopy, for each plane that is imaged in the sample the entire volume is illuminated, and sectioning is achieved by rejecting out-of-focus fluorescent emission with a confocal pinhole. In contrast, in SPIM only the in-focus plane of the sample is illuminated, so no out-of-focus light is generated and all of the emitted fluorescence can be used. This means that SPIM provides more information per unit of exposure of the sample. One can therefore achieve higher signal-to-noise ratios in SPIM than would be possible in confocal imaging without excessive photo-bleaching or damage. Similarly, because of the lower photo-toxicity, with SPIM one can image more often and over longer periods of time.

The following steps assume a custom-built SPIM instrument similar to that described in Greger et al. (2007). Although SPIM induces substantially less photo-damage than confocal imaging, it is still advisable to minimize over-exposure of the sample. One typically begins with the camera gain set to maximum and the laser power ∼10% of maximum. If the resulting signal-tonoise ratio is insufficient, the gain can be reduced and/or the laser power increased. Before imaging, align the SPIM so that the light sheet coincides with the focal plane of the detection arm, and adjust the light sheet so that it is as narrow as possible, but of uniform thickness over the camera's field of view. In our custom-built SPIM, this is done by adjusting a gimble mirror and adjustable slit that were built into the illumination path for this purpose. However, the procedure will vary between different implementations of SPIMs. Insert the sample with the region of interest to be imaged facing approximately toward the detection objective lens. See Swoger et al. (2011) for a discussion of how to optimally orient the sample. Using transmission contrast, try to center the approximate region of interest in the field of view of the camera. Depending on the developmental stage of the neuromasts, this may not be possible in transmission, but at a minimum it should be possible to find the myoseptum. This will save time and exposure of the sample when working in fluorescence. Switching to fluorescence contrast, use the sample translation stage to locate and center a neuromast in the field of view. Determine the limits of the axial scanning as described for confocal imaging. Typical slice spacing is 1–2μm. Save the positioning parameters to define the scan for that neuromast. Translate the embryo to locate another neuromast and repeat these procedures until about 4 scans have been defined. The exact number that can be done will depend on the number of slices in each stack and the maximum time between repeated stack acquisitions. Set parameters for each desired channel: emission filter, laser wavelength and intensity, and camera gain and exposure. Multiple channels can be acquired in plane-interlaced mode. The parameters for each channel should be selected to optimize image quality while imaging a typical neuromast "live." Set the timing parameters: temporal spacing of the sequential acquisitions, *t*, (typically 2–5 min) and total time of the experiment (anywhere from minutes to days, as desired). *t* will be limited by the time it takes to scan each neuromast and the number of neuromasts scanned. Before beginning the full time-lapse, it can be helpful to do a single scan of all of the neuromasts, in order to determine the minimum value of *t*. If this turns out to be longer than the desired *t* it will be necessary to reduce the required time by, e.g., reducing the number of neuromasts imaged. See also the "The acquisition time is too long" section under Troubleshooting, below. Choose a filename prefix and destination folder, and start the time-lapse acquisition.

During the initial 2 or 3 h of the scanning, occasionally check that the neuromasts are each still centered in their fields of view. Because the agarose is not a completely rigid gel it can relax slightly when the sample is first put into the microscope, resulting in a shift of the region(s) of interest. This is especially important when imaging freshly deposited neuromasts, because these can still move relative to the surrounding tissue even if the agarose is stable. Any drifts can be corrected "onthe-fly" by updating the position parameters in the software user interface.

#### **DATA PROCESSING**

Open the TIFF series in ImageJ/Fiji. Convert stack to Image5D using the Image5D Plugin. When requested use "Z" for 3rd dimension and "Time" for 4th dimension. Fill in the details of the time lapse according to your log book. Project the Z-stack to a single plane using the project Z-stack option from the Image5D plugin. To add a time counter, use the Time stamper plugin. Fill in the details of the time lapse according to your log book. Save the file as a TIFF file. To make a movie save the file as AVI and choose an appropriate frame rate for your needs. As a reference a 12 h long time lapse with an acquisition time of 2 min takes 36 s to play with a speed of 10 frames per second. The timing of data processing ranges from ∼15 min to 6 h per dataset.

#### **IF PROCESSING A DATA SET OBTAINED USING** *Tg[Cldnb:mGFP; SqET4]* **WITH THE PURPOSE OF CELL TRACKING**

Open the TIFF series in ImageJ/Fiji and reverse the stack using the reverse tool in Image/Stacks/Tools. Save the LIF file as Image Sequence in a designated folder with the name "STACKNAME\_Reversed STACK TIFFS." It is crucial to tick the option use slice labels as file names. You have just inverted the stack so that the last time point of the acquired data is now the first to be displayed and the first time point is the last to be displayed. CRUCIAL! Remove the scale of the stack in Analyze/Set Scale. Convert stack to Image5D using the Image5D Plugin. When requested, use "Z" for 3rd dimension and "Time" for 4th dimension. Fill in the details of the time lapse according to your log book. Open the MtrackJ plugin. In the displaying menu tick the option "Display only track points at current time" and reduce the "track width" to 0. Identify the cell(s) to be tracked. Press ADD in the MtrackJ panel and with the mouse cursor over the cell of interest press the mouse button. The program will save the details (XYZT coordinates) of the selection and move to the next time index after adding the point. If the selection is no longer visible on the current Z-axis use the Z-axis scroll bar in the Image5D window to relocate the region of interest. Repeat these steps until the end of the time lapse. Save the tracking using the saving function In the MtrackJ control panel. In the MtrackJ control panel press the Measure Button. Observe that the measurement window shows for each time point the details of the selection (XYZT coordinates). Save the measurement results as an excel file. If more cells will be tracked, restart MtrackJ to clear the memory, go to the first image of the stack and identify the cell to be tracked. Repeat the described procedure for each desired tracking.

Open the folder "STACKNAME\_Reversed STACK TIFFS" where the stacks were saved as an Image sequence. Observe that each file name contains a reference to the time point and a Z coordinate. Using the details of the selection (XYZT coordinates) in the excel file select for each time point the file corresponding to the Z coordinate of the selection. If tracking more than one cell use only the Z coordinates of one of the cells for the selection. The selected files should be copied to a new folder named "STACKNAME\_Z\_SELECTION." You have just created a time lapse of the dataset that follows the cell(s) of interest. Open the files in "STACKNAME\_Z\_SELECTION" as image sequence in ImageJ. Open MtrackJ. Load the previous saved tracking results. To show the tracks along all time points, in the MtrackJ control panel Displaying options select a Track width size of 3 pixels. If do not want the number of each MtrackJ point to be shown in the MtrackJ control panel under the section Displaying/Font size choose 0 pixels. Crop the time lapse to the region of interest. For this use the rectangular selection tool to define the region of interest and then use the function crop in Image/Crop. Reverse the reversed time lapse using the reverse tool in Image/Stacks/Tools. You have just inverted the time lapse so that the first time point of the acquired data is now the first to be displayed and the last time point is the last to be displayed. To add a time counter, use the Time stamper plugin. Fill in the details of the time lapse according to your log book. This is the final TIFF file. Save it with the appropriate name. To make a movie from the file save it as an AVI and choose an appropriate frame rate for your needs. As a reference a 12 h long time lapse with an acquisition time of 2 min takes 36 s to play with a speed of 10 frames per second (**Figures 3**, **6**).

#### **TROUBLESHOOTING**

#### **THE ACQUISITION TIME IS TOO LONG**

The acquisition time can be reduced by:

#### **THE FISH DIE DURING IMAGE ACQUISITION**

#### **THE DATA SET DOES NOT OPEN IN IMAGE J**