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. 2025 Jun 28;63(4):e70021. doi: 10.1002/dvg.70021

Tracing Early Migratory Neurons in the Developing Nose Using Contactin‐2 (Cntn2) CreERT2

Enrico Amato Jr 1, Alexis M Semon 1, Paolo E Forni 1,✉
PMCID: PMC12205475  PMID: 40580015

ABSTRACT

Neuronal migration during embryonic development is a fundamental process. In the developing nose of rodents, neurons that form during early neurogenic waves in the olfactory placode leave this structure to migrate toward or into the developing brain as part of the migratory mass. This mass includes gonadotropin‐releasing hormone‐1 (GnRH‐1) neurons, pioneer/terminal nerve (TN) neurons, as well as neural crest‐derived olfactory glial cells called olfactory ensheathing cells. There have been a limited number of molecular markers available to effectively trace and functionally manipulate the early migratory neurons that originate in the olfactory region. Contactin‐2 (Cntn2), also known as transiently expressed axonal surface glycoprotein‐1 (TAG‐1), has been used to label various developing neuronal populations, including the commissural neurons of the spinal cord, motor neurons, and TN neurons. Previous single‐cell RNA sequencing analyses of the developing olfactory system have identified Cntn2 expression in the TN, suggesting that Cntn2 is a suitable molecular marker for studying nasal migratory neurons. To trace Cntn2 expression in the developing olfactory system, we generated an inducible Cntn2CreERT2 mouse line. In this study, we outline how this mouse line can serve as an effective tool for time‐controlled chimeric manipulation of specific neuronal populations of interest.

Keywords: Contactin‐2 (Cntn2), GnRH neurons, migratory mass, olfactory development, spinal cord, terminal nerve

1. Introduction

Contactin‐2 (Cntn2), also known as transiently expressed axonal surface glycoprotein‐1 (Tag1), belongs to the immunoglobulin (Ig) superfamily (Furley et al. 1990). Cntn2 has been implicated in various developmental processes such as cell adhesion (Gurung et al. 2018), axonal coalescence (Wolman et al. 2008), neurite outgrowth, axon pathfinding, and regulation of neuronal migration (Suter and Jaworski 2019). Cntn2 can facilitate cell–cell contacts through interactions with other cell adhesion molecules and extracellular matrix molecules (Fan et al. 2024).

Besides its specific biological functions, Cntn2 has been widely used as a developmental stage‐specific molecular marker for various neuronal populations, including commissural neurons of the spinal cord, motor neurons, and putative terminal nerve (TN) neurons in the nose of rodents (Amato Jr. et al. 2024; Dodd et al. 1988; Taroc et al. 2017; Tessier‐Lavigne et al. 1988; Yamamoto et al. 1986).

The GnRH‐1 neurons are a hypothalamic neuronal population that produces and releases the decapeptide GnRH‐1 (Schwanzel‐Fukuda and Pfaff 1989; Wray 2001; Wray et al. 1989). The pulsatile release of GnRH‐1 regulates the hypothalamic–pituitary‐gonadal hormonal axis, which is responsible for reproductive development and hormone regulation in adulthood (Kaprara and Huhtaniemi 2018; Pohl and Knobil 1982; Schwanzel‐Fukuda et al. 1989). During embryonic development, the GnRH‐1 neurons migrate from the olfactory pit to the hypothalamus as part of a migratory mass that also includes pioneer/TN neurons and olfactory glial cells called olfactory ensheathing cells (Barraud et al. 2010; Barraud et al. 2013; Forni, Taylor‐Burds, et al. 2011; Taroc, Naik, et al. 2020).

The migration of GnRH‐1 neurons has been a subject of investigation for several decades (Duittoz et al. 2022; Schwarting et al. 2007). However, the lack of molecular markers and precise genetic entry points to study, follow and manipulate the early neurons forming in the olfactory area has been a significant limitation.

While for decades, this migration was described to occur along the axons of vomeronasal or olfactory neurons (Schwarting et al. 2001; Yoshida et al. 1995), other studies suggested that the GnRH‐1 neurons invade the brain on the axons of pioneer neurons, forming the putative TN (Amato Jr. et al. 2024; Demski and Schwanzel‐Fukuda 1987; Schwanzel‐Fukuda et al. 1987; Schwanzel‐Fukuda and Pfaff 1989; Taroc, Katreddi et al. 2020; Taroc et al. 2017).

Peripherin (Jennes 1992; Wray et al. 1994) and Cntn2 (Schwarting et al. 2001; Schwarting et al. 2004) have been used as markers to follow the development of the GnRH‐1 migratory track (Schwarting et al. 2001). Cntn2 immunoreactivity is notably stronger in axons than in cell bodies, making it challenging to identify which cells express it. However, the question of which cells in the migratory mass express Cntn2 (Taroc et al. 2017) has never been clearly addressed because of the transient expression of this gene and the intermingling of axons and cell bodies within the migratory mass (Chao et al. 2009).

The migratory neurons that form in the olfactory placode are among the first to express neuronal markers (Fornaro et al. 2007; Gong and Shipley 1995). In a recent study, Prokr2, a gene associated with defective GnRH‐1 migration to the brain, and the Microtubule‐associated protein 2 (MAP2) have been respectively identified as an early genetic marker and antigen for the presumptive TN neurons in the developing nose (Amato Jr. et al. 2024). Notably, Prokr2Cre tracing allowed for the identification of Prokr2‐expressing TN cells in the early developing nose as a distinct cell population, separate from GnRH‐1 immunoreactive neurons and early olfactory and vomeronasal sensory neurons. Single‐cell sequencing of the Prokr2‐expressing TN cells indicated that these cells are enriched for Peripherin and Cntn2/Tag‐1.

Cntn2 loss of function can compromise the development of the nervous system (Suter et al. 2020). So, to further characterize the TN and test if Cntn2 can be used as a timely controlled genetic entry point in the developing GnRH‐1‐TN system, we designed a new line of inducible Cntn2CreERT2 mice. The P2A auto cleaving strategy was adopted to faithfully drive Cre expression, relying on endogenous cis‐regulatory elements (Figure 1a), without interfering with Cntn2 endogenous expression and function (Donnelly, Hughes, et al. 2001; Donnelly, Luke, et al. 2001; Tang et al. 2009). In this article, we describe how this new Cntn2CreERT2 mouse line can serve as a valuable tool for chimeric manipulation and neuron tracing in a timely and controlled fashion.

FIGURE 1.

FIGURE 1

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Strategy for generation of Cntn2CreERT2 mice and expression of Cntn2 in the Spinal Cord. (a) Construct used in the creation of Cntn2CreERT2 mice. Based on strategy report provided by Cyagen. (b,b′) Parasagittal sections of E11.5 and E14.5 C57BL/6J WT embryos immunostained for Cntn2, highlighting expression in the dorsal root ganglia (DRG). (c) In the transverse section of an E13.5 embryo, Cntn2 is expressed by the DRG and within the commissural neurons (notched arrowhead), passing through the floorplate (FP) of the neural tube (NT). Scale bars in (b, b′, c), 100 μm.

2. Results

2.1. Cntn2CreERT2 Recombination Faithfully Recapitulates Cntn2 Expression in the Spinal Cord and Dorsal Root Ganglia

During development, Cntn2 is transiently expressed by subsets of spinal cord commissural neurons and by neurons in the dorsal root ganglia (Furley et al. 1990; Karagogeos et al. 1991) (Figure 1b,c). We tested Cntn2CreERT2 mediated recombination by injecting tamoxifen at E11.5 and collecting Cntn2CreERT2 +/−/R26tdTomato; Ai14+/− embryos one day post injection (1 DPI) at E12.5 (Figure 2b).

FIGURE 2.

FIGURE 2

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Cntn2CreERT2 recombination in the spinal cord overlaps with Cntn2 immunoreactivity. (a) Immunofluorescence for Cntn2 (Contactin 2, green) on transverse E12.5 C57 BL/6J WT tissue reveals the dorsal root ganglion (DRG) proximal to the neural tube (NT) are positive. The commissural neurons are highlighted by Cntn2 immunostaining, passing through the floor plate (FP) (notched arrowhead). (b) Cntn2CreERT2 +/−/Ai14+/− mice were injected with tamoxifen at E11.5 and collected at E12.5. (c‐c”, d‐d”) Double Immunofluorescent staining against Cntn2 (green) and tdTomato (magenta) on parasagittal and transverse E12.5 Cntn2CreERT2 +/−/Ai14+/− animals highlights traced neurons immunopositive for Cntn2 in the DRGs (arrowheads). Scale bars in (a, c, d) 100 μm; (c′, c″, d′, d″), 50 μm.

Cntn2 is known to have a transient expression; therefore, some of the cells that undergo Cntn2CreERT2 mediated recombination are immunoreactive for Cntn2 one or 2 days after recombination, while others will not be, as Cntn2 levels decrease. In line with this, immunofluorescent staining against Cntn2 and the reporter gene tdTomato revealed a faithful expression pattern of the reporter gene, overlapping with Cntn2 immunoreactivity in 75% (SD ± 4%; n = 3) of all the cells that underwent tamoxifen‐induced Cre recombination 1 day before (Figure 2c,d). We observed that Cntn2 tracing is specific to areas in the spinal cord where Cntn2 expression is found (commissural neurons and dorsal root ganglia); this shows that the Cntn2CreERT2 mouse model accurately traces cells that express Cntn2.

2.2. Cntn2 is Expressed by the Neurons of the Nasal Migratory Mass

Cntn2 has been extensively used to follow the trajectory of migrating GnRH‐1 neurons (Duittoz et al. 2022; Schwarting et al. 2001; Taroc et al. 2017; Yoshida et al. 1995).

In a recent study, we traced the pioneer/TN neurons forming from the olfactory placode, showing that these cells are genetically distinct from the olfactory, vomeronasal, and GnRH‐1 neurons (Amato Jr. et al. 2024). Gene expression profiling of the putative TN cells highlighted the enrichment of Peripherin, Robo3, and Cntn2 expression, which have been previously suggested to mark the GnRH‐1 migratory scaffold (Taroc et al. 2019; Taroc et al. 2017) (Figure 5a). Immunohistochemistry against Cntn2 on control animals from E10.5 to E13.5 confirmed the Cntn2 immune detectability in axons and in sparse cell bodies in the developing olfactory area (Figure 3).

FIGURE 5.

FIGURE 5

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Contactin‐2 (Cntn2) as a genetic entry point to manipulate the terminal nerve: (a) Volcano plot comparing enriched genes of the sorted migratory pioneer/TN neurons with all unsorted neurons. (b–d) Developmental time‐course of Cntn2 immunoreactivity in the developing vomeronasal organ (VNO) and migratory TN from E11.5‐E13.5 using Prokr2Cre+/−/Ai14+/− mice. Immunostaining for Cntn2 (green) and Prokr2 lineage tracing (magenta) was performed at all three developmental stages. (b–b‴) At E11.5, Cntn2 immunoreactivity is in Prokr2 traced pioneer/TN neurons (arrowheads) emerging from the olfactory epithelium (OE) proximal to the forebrain (FB). (c–c‴‴) Cntn2 immunoreactivity is detected in the traced Prokr2 cells at E12.5, migrating from the VNO to the forebrain junction (FBJ). (d–d‴‴) Visualizing at E13.5, an increased number of Prokr2 traced neurons positive for Cntn2 are present in the VNO and FBJ. Scale bars in (c‴′–c‴‴, d‴′‐d‴‴) 25 μm; (b′–b‴, c′–c‴, d′–d‴) 50 μm; b–d 100 μm.

FIGURE 3.

FIGURE 3

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Contactin‐2 (Cntn2) is expressed by neurons of the nasal migratory mass. (a–d) Developmental time‐course of immunohistochemistry against Cntn2 on C57BL/6J WT mice. (a,a′) At E10.5, Cntn2 expression is shown in the migratory pioneer/TN neurons within the migratory mass (MM) emerging from the olfactory placode (OP) and proximal to the developing forebrain (FB) indicated by the black arrowheads. (b,b′) Cntn2 expression at E11.5 is still present in the pioneer/TN neurons migrating away from the olfactory epithelium (OE). (c,c′) At E12.5, Cntn2 expression is seen in a select group of neurons, labelling cell bodies and fibers outside of the vomeronasal organ (VNO) and adjacent to the forebrain junction (FBJ). (d,d′) Observing Cntn2 expression at E13.5 shows neurons leaving the VNO and projecting toward the brain, passing the olfactory bulb (OB). Scale bars in (a′, b′) 25 μm; (c′, d′) 50 μm; (a, b) 100 μm; (c, d) 250 μm.

Cntn2 expression was found in migratory pioneer/TN neurons at all stages of development analyzed. At E12.5 and E13.5, Cntn2+ cell bodies and fibers could be observed both outside the VNO and proximal to the forebrain junction (FBJ) (Figure 3c,d).

Previous birth‐dating experiments suggested that most of the neurons that become postmitotic between E10.5 and E11.5 migrate out of the developing olfactory system (Fornaro et al. 2003; Forni, Fornaro, et al. 2011; Taroc, Naik, et al. 2020; Wray and Hoffman 1986), this suggests that the pioneer/TN neurons, like the GnRH‐1 neurons, are among the very first neurons to form in the olfactory area.

By analyzing GnRH‐1 and Cntn2 immunoreactivity in the developing olfactory system at E11.5, E12.5, and E13.5, we observed very sparse Cntn2 immunoreactivity across the cells in the putative vomeronasal organ (Figure 4), where both GnRH‐1 and TN neurons form (Amato Jr. et al. 2024; Forni et al. 2013).

FIGURE 4.

FIGURE 4

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Transient immunoreactivity of Cntn2 in the GnRH‐1 neurons within the developing olfactory system. (a–c) Developmental time‐course of GnRH‐1 (green, arrowheads) and Cntn2 (magenta, arrows) immunoreactivity in the developing olfactory system at E11.5, E12, and E13.5. (a, a′) At E11.5, three distinct populations of neurons can be observed, GnRH‐1 neurons negative for Cntn2, Cntn2 neurons negative for GnRH‐1 and GnRH‐1 neurons that express Cntn2 (notched arrowheads). The three distinct populations of neurons are not migrating and are within the vomeronasal organ (VNO). (b, b′) Visualizing at E12.5, the three groups of neurons migrate out of the VNO past the olfactory epithelium (OE) and toward the olfactory bulb (OB). The fibers of the terminal nerve (TN) are immunoreactive for Cntn2. (c,c′) At E13.5, the majority of migratory neurons have left the VNO and have now invaded the brain. The Cntn2+ TN can be observed projecting into the basal forebrain (BFB). Scale bars in (a′–c′) 50 μm; (a,c) 100 μm.

Based on Cntn2 and GnRH‐1 immunoreactivity, we identified three populations: cells only positive for Cntn2, cells reactive for both Cntn2 and GnRH‐1, and cells only positive for GnRH1. Specifically, we found that at E11.5, only 15% (SD ± 18%; n = 3) of the GnRH‐1 immuno‐positive neurons expressed detectable Cntn2, 45% (SD ± 3%; n = 3) at E12.5, and 39% (SD ± 3%; n = 3) at E13.5 (Figure 4a–c). These data suggest asynchronous and transient expression of Cntn2 across the GnRH‐1 neurons.

2.3. Cntn2 Can Be Used as a Genetic Entry Point to Manipulate the Cells of the Nasal Migratory Mass

Prokr2Cre lineage tracing and Map2 immunoreactivity can be used to visualize the cell bodies of the developing TN (Amato Jr. et al. 2024). Previously acquired single‐cell RNA sequencing data at E14.4, indicate Cntn2 is enriched in the TN neurons compared to other neurons in the developing olfactory system, (Figure 5a) (Amato Jr. et al. 2024). Based on this, we decided to monitor Cntn2 immunoreactivity in the migratory TN of Prokr2Cre traced animals at various developmental stages. We characterized Cntn2 expression in Prokr2 traced cells using Prokr2Cre+/−/Ai14+/− mice at E11.5‐E13.5 (Figure 5b–d).

At E11.5, Cntn2 immunoreactivity was found in the majority (~70% SD ± 12%; n = 3) of the Prokr2 traced pioneer/TN neurons emerging from the developing olfactory/vomeronasal epithelium (OE/VNE). However, as Cntn2 has a transient expression, its immunoreactivity dropped to 58% (SD ± 1%; n = 3) at E12.5 and to 36% (SD ± 7%; n = 3) of the Prokr2 traced cells at E13.5. Based on these observations, we identified E11.5 as an attractive time point to induce the Cntn2‐Cre‐driven genetic tracing in the migratory neurons forming the presumptive TN.

2.4. Using Cntn2CreERT2 Tracing to Follow the Pioneer Neurons/Terminal Nerve

Cntn2CreERT2+/− mice were mated with Ai14+/− reporter mice. Pregnant dams were injected with tamoxifen (75 mg/kg body weight) at E11.5.

We analyzed the Cntn2 tracing at E12.5 (1‐day post‐injection [DPI]), E13.5 (2 DPI), and E15.5 (4 DPI). We combined Cntn2 tracing with Cntn2 immunostaining (Figure 6a–c). We quantified the percentage of Cntn2‐traced cells displaying Cntn2‐immunoreactive bodies (Figure 6d). We found Cntn2 tracing and immunoreactivity overlapping in 17% (SD ± 8%; n = 3) of the cells at E12.5, 38% (SD ± 1%; n = 3) at E13.5 and in 21% (SD ± 4%; n = 3) of the cells at E15.5, providing more evidence for Cntn2's transient expression pattern. Cntn2 tracing was observed to label many migratory TN cells within the nasal area at E12.5 and E13.5. Notably we observed a larger number of cells over time with a significant increase at E13.5 (Figure 6e, p = 0.0002). However, from E13.5 to E15.5, a significant decrease in the number of traced cell bodies was observed in the nose, suggesting that a portion of the traced neurons might have either died or migrated into the brain (Figure 6e, p = 0.0077). This prompted us to investigate Cntnt2Cre recombination in the GnRH‐1 neurons.

FIGURE 6.

FIGURE 6

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Using Cntn2CreERT2 to label cell bodies and Cntn2 to highlight the axons of pioneer neurons/terminal nerve (TN). (a–d) Developmental time‐course of Cntn2 immunoreactivity in the developing vomeronasal organ (VNO) and migratory TN from E12.5 (1 DPI), E13.5 (2 DPI), and E15.5 (4 DPI) using Cntn2CreERT2 +/−/Ai14+/− mice. Immunostaining for Cntn2 (green) and Cntn2 lineage tracing (magenta) was performed at all three developmental stages. Arrowheads indicate neurons that are traced for Cntn2, and empty arrowheads indicate neurons that are both traced for Cntn2 and express Cntn2. (a–a‴) E12.5 reveals Cntn2 traced pioneer/TN neurons labelling cell bodies and Cntn2 immunoreactivity within the axons and fibers. (b–b‴) At E13.5, Cntn2 immunoreactivity was found to be greater in Cntn2‐traced neurons migrating out of the VNO, toward the forebrain junction (FBJ) proximal to the olfactory bulb (OB). (c–c‴) At E15.5, Cntn2 immunoreactivity was not detected in the Cntn2‐traced neurons near the VNO. Red blood cells are marked with asterisks. (d) Quantifications of Cntn2 tracing and traced neurons that are positive for Cntn2 expression from E12.5‐E15.5, show an increase in Cntn2 expression at E13.5. Mean for 1DPI: Cntn2 tracing, 83% (SD ± 8%; n = 3); Cntn2 tracing+Cntn2 expression, 17% (SD ± 8%; n = 3); mean for 2DPI: Cntn2 tracing, 62% (SD ± 1%; n = 3); Cntn2 tracing + Cntn2 expression, 38% (S.D. ± 1%; n = 3); mean for 3DPI: Cntn2 tracing, 80% (SD ± 4%; n = 3); Cntn2 tracing+Cntn2 expression, 21% (SD ± 4%; n = 3). Individual percentage values are represented as dots. (e) Quantification of Cntn2 traced neurons in the nasal area from E12.5‐E15.5 shows significant changes over time (±SD, one‐way ANOVA, p < 0.05, F = 40.07, n = 3, top asterisk), multiple comparisons significance was determined via Dunnett's Test (***p = 0.0003). The data used for the one‐way ANOVA had normal distribution, equal variance and was adjusted for multiple comparisons (Mean for 1DPI, 18% (S.D. ± 6%; n = 3); 2 DPI, 49% (S.D. ± 1%; n = 3); 3 DPI 34% (S.D. ± 4%; n = 3). Individual average values are represented as dots. Scale bars in a′–a‴, b′–b‴, c′–c‴, 25 μm; (a, b, c) 100 μm.

We found no spontaneous recombination in non‐injected Cntn2CreERT2+/−/Ai14+/− (Figure S1).

2.5. GnRH‐1 Neurons Are Positive for Cntn2 Tracing

To understand if a portion of the Cntn2 traced migratory neurons are the GnRH‐1 neurons, we performed immunostaining against GnRH‐1 on Cntn2CreERT2+/− traced animals at 1,2 and 4 DPI (tamoxifen injection at E11.5). Both Cntn2 traced neurons and traced neurons also expressing GnRH‐1 were observed in the migratory mass together with GnRH‐1 neurons negative for tracing (Figure 7). Interestingly, we observed after one tamoxifen injection at E11.5, the number of traced cells immunoreactive for GnRH‐1 increased over time from 30% (SD ± 7%; n = 3) at 1 DPI to 58% (SD ± 25%; n = 3) at 4 DPI (Figure 7a–d), suggesting that Cntn2 expression and therefore recombination precedes GnRH‐1 immune detectability. Notably, the Cntn2 recombination in GnRH‐1 neurons was higher than what we expected based on Cntn2 immunoreactivity in the cell bodies at E11.5 (15%). Cntn2 tracing+ TN projections into the basal forebrain (bFB) were observed at E15.5 proximal to GnRH‐1 immunoreactive fibers (Figure 7e).

FIGURE 7.

FIGURE 7

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The GnRH‐1 neurons are positive for Contactin‐2 (Cntn2) tracing. (a–e‴) Double immunostaining of GnRH‐1 (green, arrows) with Cntn2 tracing (magenta, arrowheads) and GnRH‐1 neurons that are traced for Cntn2 (white, empty arrowheads) in a time course from E12.5 (1 day post‐injection (DPI)), 2 DPI E13.5 and 4 DPI E15.5. (b) Cntn2CreERT2 +/−/Ai14+/− mice were injected with tamoxifen at E11.5 and collected at E12.5‐E15.5. The relationship between GnRH‐1 neurons (green) and GnRH‐1 neurons that are traced (white) is depicted by the pie charts next to the images. (a–a‴) E12.5 shows Cntn2 traced neurons, GnRH‐1 neurons, and Cntn2 traced neurons positive for GnRH‐1 expression within and outside of the vomeronasal organ (VNO); (c–c‴) At E13.5 there is an increase of the three populations migrating out of the VNO; Cntn2 traced neurons can be seen in the olfactory epithelium (OE) and olfactory bulb (OB). Red blood cells are marked with asterisks. (d–d‴) E15.5 shows all three populations of neurons invading the brain at the forebrain junction (FBJ), while very few are still migrating out from the VNO. Red blood cells are marked with asterisks. (e–e‴) A magnification of the basal forebrain (bFB) at E15.5, with the three populations of neurons invading. Scale bars in (c′–c‴ 25 μm; a′–a‴, d′–d‴ 50 μm; (a, c, d, e) 100 μm.

The GnRH‐1 neurons become post‐mitotic in a non‐synchronous fashion (Jasoni et al. 2009). To further evaluate Cntn2 tracing and expression in GnRH‐1 neurons, we injected Cntn2CreERT2 +/− animals at E11.5 and E12.5, collecting at E15.5, a stage at which the GnRH‐1 neurons invade the brain extensively (Figure 8). From this experimental paradigm, we found a similar number of GnRH‐1 neurons were traced (51% SD ± 16%; n = 3), comparable to one injection at E11.5.

FIGURE 8.

FIGURE 8

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Double injections at E11.5 and E12.5 trace GnRH‐1 neurons invading the brain. (a) Cntn2CreERT2 +/−/Ai14+/− mice were injected with tamoxifen at E11.5 and E12.5 and collected at E15.5. (b–b‴) Double immunostaining of GnRH‐1 (green, arrows) and Cntn2 tracing (magenta, arrowheads) at E15.5. Many GnRH‐1 neurons are observed to be invading the basal forebrain (bFB) with Cntn2 traced neurons and GnRH‐1 neurons that are traced for Cntn2 (white, empty arrowheads). The olfactory bulb (OB) was positive for Cntn2 tracing. Scale bars in (b, 200 μm; (b′–b‴) 100 μm.

Notably, we observed traced cell bodies negative for GnRH and TN fibers at the forebrain junction and invading the brain, suggesting that cells of the putative TN might invade the brain together with the GnRH‐1 neurons. The transcription factor Isl1 is an early marker for maturing GnRH‐1 neurons, which is expressed by virtually all GnRH cells in mice (Taroc, Katreddi et al. 2020; Zouaghi et al. 2025). We therefore performed Isl1 immunostaining on Cntn2CreERT2+/− traced animals (Figure 9). As observed for GnRH‐1 immunoreactivity we also found that the percentage of Isl1 immunodetectable traced neurons increases over time: from 22% (SD ± 10%; n = 3) at 1 DPI to 48% (SD ± 4%; n = 3) at 2 DPI. This suggests that Cntn2 expression precedes both Isl1 and GnRH‐1 immune detectability.

FIGURE 9.

FIGURE 9

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Isl1 immunoreactivity precedes Cntn2 expression. (a–c) Double immunofluorescent staining of Isl1 (green, arrows) and Cntn2 tracing (magenta, arrowheads). (b) Cntn2CreERT2 +/−/Ai14+/− mice were injected with tamoxifen at E11.5 and collected at E12.5 and E13.5. Isl1 expression is depicted by the pie charts next to the images, Isl1+ neurons (green) and Isl1 neurons that are traced (white). (a–a‴, c–c‴) At E12.5 and E13.5, migratory neurons expressing Isl1, Cntn2 tracing, and Isl1 immunopositive traced neurons (white, empty arrowheads) were observed to be approaching the brain, proximal to the olfactory bulb (OB). Asterisks indicate red blood cells. Scale bars in (a′–a‴) 25 μm; (c′‐c‴) 50 μm; (a, c) 100 μm.

3. Discussion

Temporally controlled chimeric genetic recombination in neurons is a valuable method for studying the developmental and physiological roles of genes in specific neuronal subsets at distinct developmental stages. During the early stages of the development of the peripheral and central nervous systems, neurons share a large number of generic neuronal genes, making cell type‐specific recombination or tracing a challenging task (Fornaro et al. 2007). The olfactory placode is among the very first neurogenic areas in rodent embryos. In fact, migratory neurons can be found emerging from the invaginating olfactory placode as early as E10, which is around two days before cortical neurogenesis starts (Chen et al. 2017).

Contactin‐2 can exhibit a transient expression modality in developing neurons, which is why it was previously referred to as transient axonal glycoprotein‐1 (TAG1) (Wolfer et al. 1994; Yoshida et al. 1995). Cntn2's differential, asynchronous and transient expression across neuronal types, has made Cntn2 a useful marker to follow specific types of neurons at defined developmental stages.

In this paper, we introduced and characterized a newly generated Cntn2CreERT2 mouse line, a novel genetic tool for temporally controlled chimeric recombination in neurons. Our data show that this Cntn2‐P2A‐CreERT2 model faithfully recombines in Cntn2‐expressing neurons, such as the commissural neurons (Figures 1, 2) of the spinal cord, dorsal root ganglia (Dodd et al. 1988; Shu et al. 2022; Suter et al. 2020) as well as in the migratory pioneer/TN neurons of the developing olfactory system, including the GnRH‐1 neurons (Figures 6, 7, 8) (Ware et al. 2016; Wolfer et al. 1994; Yamamoto and Schwarting 1991). The transient and asynchronous expression of Cntn2 across neurons makes the Cntn2CreERT2 mouse line a valuable tool for neurodevelopmental studies aimed at understanding the role of specific guidance molecules or genes involved in neuronal migration (Figures 3, 4).

Cntn2 expression in the developing olfactory area has been observed in neurons believed to form the TN, which serves as the GnRH‐1 migratory scaffold (Casoni et al. 2016; Duittoz et al. 2022; Schwanzel‐Fukuda and Pfaff 1989; Taroc et al. 2017). However, given the strong immunoreactivity in the axons and weak immunoreactivity in the cell soma, along with the highly intertwined cell bodies and axons of the olfactory migratory mass, it has never been fully established whether the GnRH‐1 neurons themselves express Cntn2.

We recently discovered that Prokr2 Cre recombination selectively labels the putative TN neurons, which we described as migratory neurons of the nasal area that are neither immunoreactive for GnRH‐1 nor for the transcription factor Isl1. Notably, Isl1 is a transcription factor that is highly enriched in the GnRH‐1 neurons (Lund et al. 2020; Taroc, Katreddi et al. 2020; Zouaghi et al. 2025). Single‐cell RNA sequencing of Prokr2+ presumptive TN cells revealed that Cntn2 is a highly enriched gene in these neurons (Figure 5) (Amato Jr. et al. 2024; Martin et al. 2011; Pitteloud et al. 2007). Using Prokr2Cre recombination as a reference, we found that at E11.5, Prokr2 tracing, and Cntn2 immunoreactivity overlapped with putative TN cells in the nasal area (70%). Based on this, this stage is appropriate for tamoxifen induction of Cntn2CreERT2 recombination in the TN of mice. After tamoxifen induction at E11.5 the Cntn2CreERT2+/−/Ai14+/− embryos were analyzed at multiple intervals post injection (Figure 6). This analysis revealed recombination in the TN as well as in the GnRH‐1 neurons. After a single tamoxifen injection at E11.5, there was an increase in the number of GnRH‐1 neurons positive for immune detectability in traced cells over time (Figure 7). Isl1 immune detectability was also found to increase in the traced neurons (Figure 9). This suggests Cntn2 is an early gene expressed prior to GnRH‐1/Isl1 immunoreactivity.

Our data suggest that Cntn2 is transiently expressed in both GnRH‐1 and TN neurons. Notably at E12.5, we found that approximately half of the GnRH‐1 neurons appear to be Cntn2 immunoreactive (Figure 4). In line with this, tamoxifen injections in Cntn2CreERT2+/−/Ai14+/− at E11.5 and E12.5 traced around 50% of the GnRH‐1 neurons (Figure 8). These data indicate that Cntn2 is a common gene among GnRH‐1 neurons and other neurons in the nasal migratory mass and that Cntn2 starts to be expressed before GnRH becomes immunodetectable.

In summary, this study introduces and characterizes the Cntn2CreERT2 mouse line. This line serves as a reliable tool for accurately tracing and recombining Cntn2 expressing neurons. By leveraging the transient expression of Cntn2, this model enables precise chimeric tracing of neuronal populations, including commissural neurons, dorsal root ganglia, and migratory neurons in the developing olfactory system.

Analyzing this mouse, we confirmed extensive Cntn2 immunoreactivity (~70%) in the Prokr2‐traced TN neurons, neurons that we proposed to form the GnRH‐1 migratory scaffold. However, tamoxifen‐induced recombination at distinct developmental stages highlighted that Cntn2 is also transiently expressed in GnRH‐1 neurons, making this model a valuable genetic entry point for studying candidate genes in the early development of the olfactory and GnRH‐1 systems.

4. Materials and Methods

4.1. Animals

4.1.1. Mouse Lines

The custom mouse line C57BL/6J‐Tg(Cntn2‐creERT2)Forni was developed using CRISPR/Cas9 technology by Cyagen Biosciences, utilizing the C57BL/6J background strain. Cntn2CreERT2 was generated by targeting exon 23 on the Cntn2 gene using CRISPR/Cas9 to insert the donor vector containing the “P2A‐CreERT2” cassette (Figure 1a). The targeting vector was designed with homology arms and co‐injected into fertilized mouse eggs with the Cas9 mRNA and Cntn2 gRNA (Sequence, matching the reverse strand of the gene: AGCGTTGAGATCAGAGCCTCTGG). F0 Founder animals were identified through PCR and subsequently bred with C57BL/6J wild‐type mice to assess germline transmission and facilitate further F1 animal generation. The Cntn2CreERT2 mouse line will be available at The Jackson Laboratory Repository (JAX Stock No. 040586); https://www.jax.org/query.

R26tdTomato Ai14 (JAX stock 007914) mice were maintained on a C57BL/6J background. Cntn2CreERT2 and Ai14 genotypes were confirmed using PCR analysis using the following primers for Ai14: IMR0920: AAGGGAGCTGCAGTGGAGTA; IMR9021: CCGAAAATCTGTGGGAAGTC; IMR9103: GGCATTAAAGCAGCGTATCC; IMR9105: CTGTTCCTGTACGGCATGG. For Cntn2CreERT2 two primer sets were used interchangeably: generic Cre primers: Cre FWD: AGG TGT AGA GAA GGC ACT TAG C; Cre RVS: CTA ATC GCC ATC TTC CAG CAG G; and specific Cntn2CreERT2 primers: Cntn2 WT F4: TAAGGCCTCCATATGACTTTCCTC; Cntn2 WT R7: AAAATTCCTTGCCTGGTTCTATCC; Cntn2 Cre F7: AAGAACGTGGTGCCCCTCTAT; Cntn2 Cre R7: AAAATTCCTTGCCTGGTTCTATCC. Animal euthanasia was done using CO2, then cervical dislocation. Prokr2Cre+/− mice (Mohsen et al. 2017) were previously described (Amato Jr. et al. 2024). Experiments were performed with mice of either sex.

All animal experiments were completed in accordance with the guidelines of the Animal Care and Use Committee at the University of Albany, SUNY.

4.1.2. TAM Preparation and Treatment

Tamoxifen (Sigma–Aldrich), CAS # 10540‐29‐1, was mixed and dissolved in corn oil at a concentration of 20 μg/μL. Time‐mated females were injected intraperitoneally at the chosen embryonic day with a dosage of 75 mg/kg body weight, and embryos were collected at the desired time point. The observation of the vaginal plug from time‐mated females was defined as E0.5 to determine embryo age. No adverse effects on development were observed after tamoxifen injections. Embryos were collected alive and within the average of 6 embryos per litter, (Mitchell et al. 2024). Mutants and controls were harvested at the expected Mendelian distribution.

4.1.3. Tissue Preparation

Collected embryos were fixed in 3.7% formaldehyde/PBS at 4°C for 2 h, then submerged in 30% sucrose overnight. Embryos were embedded and frozen in O.C.T. (Tissue‐TeK) and stored at −80°C. Cryosectioning was done using a CM3050S Leica cryostat; samples were collected on Super‐frost plus slides (VWR) at 18 μm thickness for immunofluorescent staining.

4.2. Immunohistochemistry

4.2.1. Immunofluorescence

Primary antibodies and dilutions used for this study: goat‐α‐Cntn2 (1:1000, R&D Systems), chicken‐α‐RFP (1:1000, Rockland), SW rabbit‐α‐GnRH‐1 (1:6000, Susan Wray, NIH), mouse‐α‐Isl1 (1:100, DSHB) and rabbit‐α‐RFP (1:500, Rockland). Experiments using mouse‐α‐Isl1 and rabbit‐α‐RFP required antigen retrieval which was done by submerging slides in citric acid solution heated to 95°C for 15 min, then cooling for 15 min before proceeding with the standard immunofluorescence protocol. Secondary antibodies matched for the correct species were conjugated to Alexa Fluor‐488, Alexa Flour‐594, or Alexa Flour‐680 (Invitrogen and Jackson Laboratories). Counterstaining was done using 4′,6′‐diamidino‐2‐phenylindole (1:3000; Sigma‐Aldrich) and coverslips were mounted with Fluorogel l (Electron Microscopy Sciences). Confocal microscopy image acquisition was taken with a LSM 980 microscope (Zeiss). FIJI/ImageJ was used for image analysis and quantifications. Each staining presented was replicated on three different animals.

4.2.2. Chromogen‐Based Reactions

Experiments using chromogen‐based reactions were stained as described previously (Forni, Fornaro, et al. 2011). The tissue was processed using a standard avidin–biotin–horseradish peroxidase/3,3‐diaminobenzidine (DAB) procedure. Each staining was visualized using nickel (II) sulfate heptahydrate (Sigma) to intensify the DAB reaction (black) followed by a counterstain with methyl green. Brightfield images were taken on a Leica DM4000 B LED fluorescence microscope equipped with a Leica DFC310 FX 422 camera. FIJI/ImageJ software was used for further evaluation. Staining presented were replicated on three different animals.

4.3. Cell Quantifications

All quantifications were performed using the cell counter plugin within FIJI/ImageJ. Quantification of Cntn2 tracing and Cntn2 immunoreactivity in the spinal cord was performed on E12.5 Cntn2CreERT2 traced transverse sections, and the cell bodies of the dorsal root ganglia were quantified. For all quantifications in the developing nasal area, E12.5–E15.5 Cntn2CreERT2 traced parasagittal sections were used. For each immunostaining (Cntn2, GnRH‐1 and Isl1) paired with Cntn2 tracing, cell bodies in the nasal area/migratory mass were quantified. For all quantifications, the number of cells per section (~6 sections) was averaged for each animal. Raw data are available as a supplementary spreadsheet.

4.4. Statistics

Statistical analyses were conducted using Graphpad Prism 10.4.1. At least three biological replicates were used for each quantification. For three or more variables, we employed ordinary one‐way ANOVA followed by post hoc multiple comparisons using Dunnett's test. Normality was checked using the Shapiro–Wilk test, and equal variance was determined using the Brown‐Forsythe test. Significance was established with a p value ≤ 0.05.

4.5. Single‐Cell RNA Sequencing

The volcano plot was generated using R programming software with single‐cell sequencing data available through the Gene Expression Omnibus (GEO) database, under GEO accession number GSE234871 (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE234871). These were previously described (Amato Jr. et al. 2024).

Supporting information

Figure S1. Without tamoxifen injection Cntn2CreERT2 +/− have no spontaneous recombination. (a‐b) Cartoons representing the two experimental conditions, tamoxifen‐injected and non‐injected. (c‐d″) Immunofluorescent staining on injected or non‐injected E12.5 mice stained for the pan neuronal marker HuCD (green) and anti‐tdTomato antibody (magenta). HuCD‐labeled neurons in the nasal area included cells of the vomeronasal organ (VNO), olfactory epithelium (OE), and forebrain (FB). In the non‐injected animals (c‐c″), no tdTomato was detected in the neurons (empty arrowheads), while in tamoxifen‐injected animals (d‐d″), Cntn2Cre tracing (magenta) labeled neurons in the brain and nasal epithelia (white arrowheads). (g‐h″) Neurons of the trigeminal (TG) stained for HuCD (green) and Cntn2 tracing (magenta), consistently showing no Cntn2 tracing in non‐injected embryos (empty arrowheads), while extensive recombination was found in the tamoxifen‐treated embryos. Scale bars in f‐f′, 25 μm; g‐h″, 50 μm; c‐e″, 100 μm.

DVG-63-e70021-s002.tif (2.5MB, tif)

Appendix S1. Supplementary Information.

DVG-63-e70021-s001.xlsx (46.2KB, xlsx)

Acknowledgments

This work was supported by the Eunice Kennedy Shriver National Institute of Child Health and Human Development (NICHD) under Grants 2R01HD097331 (P.E.F.) and 1R01HD114827 (P.E.F.), as well as by the National Institute on Deafness and Other Communication Disorders (NIDCD) under Grant R01DC017149 (P.E.F.). The Zeiss 980 microscope at the University at Albany was funded by the Office of the Director, NIH, under Award Number S10OD028600.

Amato Jr., E. , Semon A. M., and Forni P. E.. 2025. “Tracing Early Migratory Neurons in the Developing Nose Using Contactin‐2 (Cntn2) CreERT2 .” genesis 63, no. 4: e70021. 10.1002/dvg.70021.

Funding: This work was supported by Eunice Kennedy Shriver National Institute of Child Health and Human Development (2R01HD097331 and 1R01HD114827), National Institute on Deafness and Other Communication Disorders (R01DC017149), and NIH Office of the Director (S10OD028600).

Data Availability Statement

The data that support the findings of this study are openly available in GEO at https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE234871, reference number GSE234871.

References

  1. Amato, E., Jr. , Taroc E. Z. M., and Forni P. E.. 2024. “Illuminating the Terminal Nerve: Uncovering the Link Between GnRH‐1 Neuron and Olfactory Development.” Journal of Comparative Neurology 532: e25599. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Barraud, P. , Seferiadis A. A., Tyson L. D., et al. 2010. “Neural Crest Origin of Olfactory Ensheathing Glia.” Proceedings of the National Academy of Sciences of the United States of America 107: 21040–21045. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Barraud, P. , St John J. A., Stolt C. C., Wegner M., and Baker C. V.. 2013. “Olfactory Ensheathing Glia Are Required for Embryonic Olfactory Axon Targeting and the Migration of Gonadotropin‐Releasing Hormone Neurons.” Biology Open 2: 750–759. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Casoni, F. , Malone S. A., Belle M., et al. 2016. “Development of the Neurons Controlling Fertility in Humans: New Insights From 3D Imaging and Transparent Fetal Brains.” Development 143: 3969–3981. [DOI] [PubMed] [Google Scholar]
  5. Chao, D. L. , Ma L., and Shen K.. 2009. “Transient Cell‐Cell Interactions in Neural Circuit Formation.” Nature Reviews. Neuroscience 10: 262–271. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Chen, V. S. , Morrison J. P., Southwell M. F., Foley J. F., Bolon B., and Elmore S. A.. 2017. “Histology Atlas of the Developing Prenatal and Postnatal Mouse Central Nervous System, With Emphasis on Prenatal Days E7.5 to E18.5.” Toxicologic Pathology 45: 705–744. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Demski, L. S. , and Schwanzel‐Fukuda M.. 1987. “The Terminal Nerve (Nervus Terminalis): Structure, Function, and Evolution. Introduction.” Annals of the New York Academy of Sciences 519: 9–11. [DOI] [PubMed] [Google Scholar]
  8. Dodd, J. , Morton S. B., Karagogeos D., Yamamoto M., and Jessell T. M.. 1988. “Spatial Regulation of Axonal Glycoprotein Expression on Subsets of Embryonic Spinal Neurons.” Neuron 1: 105–116. [DOI] [PubMed] [Google Scholar]
  9. Donnelly, M. L. L. , Hughes L. E., Luke G., et al. 2001. “The ‘cleavage’ Activities of Foot‐and‐Mouth Disease Virus 2A Site‐Directed Mutants and Naturally Occurring ‘2A‐Like’ Sequences.” Journal of General Virology 82: 1027–1041. [DOI] [PubMed] [Google Scholar]
  10. Donnelly, M. L. L. , Luke G., Mehrotra A., et al. 2001. “Analysis of the Aphthovirus 2A/2B Polyprotein ‘cleavage’ Mechanism Indicates Not a Proteolytic Reaction, but a Novel Translational Effect: A Putative Ribosomal ‘skip’.” Journal of General Virology 82: 1013–1025. [DOI] [PubMed] [Google Scholar]
  11. Duittoz, A. H. , Forni P. E., Giacobini P., et al. 2022. “Development of the Gonadotropin‐Releasing Hormone System.” Journal of Neuroendocrinology 34, no. 5: e13087. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Fan, S. , Liu J., Chofflet N., et al. 2024. “Molecular Mechanism of Contactin 2 Homophilic Interaction.” Structure 32: 1652–1658. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Fornaro, M. , Geuna S., Fasolo A., and Giacobini‐Robecchi M. G.. 2003. “HuC/D Confocal Imaging Points to Olfactory Migratory Cells as the First Cell Population That Expresses a Post‐Mitotic Neuronal Phenotype in the Chick Embryo.” Neuroscience 122: 123–128. [DOI] [PubMed] [Google Scholar]
  14. Fornaro, M. , Raimondo S., Lee J. M., and Giacobini‐Robecchi M. G.. 2007. “Neuron‐Specific Hu Proteins Sub‐Cellular Localization in Primary Sensory Neurons.” Annals of Anatomy 189: 223–228. [DOI] [PubMed] [Google Scholar]
  15. Forni, P. E. , Bharti K., Flannery E. M., Shimogori T., and Wray S.. 2013. “The Indirect Role of Fibroblast Growth Factor‐8 in Defining Neurogenic Niches of the Olfactory/GnRH Systems.” Journal of Neuroscience 33: 19620–19634. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Forni, P. E. , Fornaro M., Guenette S., and Wray S.. 2011. “A Role for FE65 in Controlling GnRH‐1 Neurogenesis.” Journal of Neuroscience 31: 480–491. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Forni, P. E. , Taylor‐Burds C., Melvin V. S., Williams T., and Wray S.. 2011. “Neural Crest and Ectodermal Cells Intermix in the Nasal Placode to Give Rise to GnRH‐1 Neurons, Sensory Neurons, and Olfactory Ensheathing Cells.” Journal of Neuroscience 31: 6915–6927. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Furley, A. J. , Morton S. B., Manalo D., Karagogeos D., Dodd J., and Jessell T. M.. 1990. “The Axonal Glycoprotein TAG‐1 Is an Immunoglobulin Superfamily Member With Neurite Outgrowth‐Promoting Activity.” Cell 61: 157–170. [DOI] [PubMed] [Google Scholar]
  19. Gong, Q. , and Shipley M. T.. 1995. “Evidence That Pioneer Olfactory Axons Regulate Telencephalon Cell Cycle Kinetics to Induce the Formation of the Olfactory Bulb.” Neuron 14: 91–101. [DOI] [PubMed] [Google Scholar]
  20. Gurung, S. , Asante E., Hummel D., et al. 2018. “Distinct Roles for the Cell Adhesion Molecule Contactin2 in the Development and Function of Neural Circuits in Zebrafish.” Mechanisms of Development 152: 1–12. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Jasoni, C. L. , Porteous R. W., and Herbison A. E.. 2009. “Anatomical Location of Mature GnRH Neurons Corresponds With Their Birthdate in the Developing Mouse.” Developmental Dynamics 238: 524–531. [DOI] [PubMed] [Google Scholar]
  22. Jennes, L. 1992. “Selective Expression of Peripherin in Gonadotropin‐Releasing Hormone‐Synthesizing Neurons of the Rat.” Molecular and Cellular Neurosciences 3: 571–577. [DOI] [PubMed] [Google Scholar]
  23. Kaprara, A. , and Huhtaniemi I. T.. 2018. “The Hypothalamus‐Pituitary‐Gonad Axis: Tales of Mice and Men.” Metabolism 86: 3–17. [DOI] [PubMed] [Google Scholar]
  24. Karagogeos, D. , Morton S. B., Casano F., Dodd J., and Jessell T. M.. 1991. “Developmental Expression of the Axonal Glycoprotein TAG‐1: Differential Regulation by Central and Peripheral Neurons In Vitro.” Development 112: 51–67. [DOI] [PubMed] [Google Scholar]
  25. Lund, C. , Yellapragada V., Vuoristo S., et al. 2020. “Characterization of the Human GnRH Neuron Developmental Transcriptome Using a GNRH1‐TdTomato Reporter Line in Human Pluripotent Stem Cells.” Disease Models & Mechanisms 13: dmm040105. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Martin, C. , Balasubramanian R., Dwyer A. A., et al. 2011. “The Role of the Prokineticin 2 Pathway in Human Reproduction: Evidence From the Study of Human and Murine Gene Mutations.” Endocrine Reviews 32: 225–246. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Mitchell, T. A. , Lin J. M., Hicks S. M., James J. R., Rangan P., and Forni P. E.. 2024. “Loss of Function of Male‐Specific Lethal 3 (Msl3) Does Not Affect Spermatogenesis in Rodents.” Developmental Dynamics 253: 453–466. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Mohsen, Z. , Sim H., Garcia‐Galiano D., et al. 2017. “Sexually Dimorphic Distribution of Prokr2 Neurons Revealed by the Prokr2‐Cre Mouse Model.” Brain Structure & Function 222: 4111–4129. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Pitteloud, N. , Zhang C., Pignatelli D., et al. 2007. “Loss‐Of‐Function Mutation in the Prokineticin 2 Gene Causes Kallmann Syndrome and Normosmic Idiopathic Hypogonadotropic Hypogonadism.” Proceedings of the National Academy of Sciences of the United States of America 104: 17447–17452. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Pohl, C. R. , and Knobil E.. 1982. “The Role of the Central Nervous System in the Control of Ovarian Function in Higher Primates.” Annual Review of Physiology 44: 583–593. [DOI] [PubMed] [Google Scholar]
  31. Schwanzel‐Fukuda, M. , Bick D., and Pfaff D. W.. 1989. “Luteinizing Hormone‐Releasing Hormone (LHRH)‐Expressing Cells Do Not Migrate Normally in an Inherited Hypogonadal (Kallmann) Syndrome.” Brain Research. Molecular Brain Research 6: 311–326. [DOI] [PubMed] [Google Scholar]
  32. Schwanzel‐Fukuda, M. , Garcia M. S., Morrell J. I., and Pfaff D. W.. 1987. “Distribution of Luteinizing Hormone‐Releasing Hormone in the Nervus Terminalis and Brain of the Mouse Detected by Immunocytochemistry.” Journal of Comparative Neurology 255: 231–244. [DOI] [PubMed] [Google Scholar]
  33. Schwanzel‐Fukuda, M. , and Pfaff D. W.. 1989. “Origin of Luteinizing Hormone‐Releasing Hormone Neurons.” Nature 338: 161–164. [DOI] [PubMed] [Google Scholar]
  34. Schwarting, G. A. , Kostek C., Bless E. P., Ahmad N., and Tobet S. A.. 2001. “Deleted in Colorectal Cancer (DCC) Regulates the Migration of Luteinizing Hormone‐Releasing Hormone Neurons to the Basal Forebrain.” Journal of Neuroscience 21: 911–919. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Schwarting, G. A. , Raitcheva D., Bless E. P., Ackerman S. L., and Tobet S.. 2004. “Netrin 1‐Mediated Chemoattraction Regulates the Migratory Pathway of LHRH Neurons.” European Journal of Neuroscience 19: 11–20. [DOI] [PubMed] [Google Scholar]
  36. Schwarting, G. A. , Wierman M. E., and Tobet S. A.. 2007. “Gonadotropin‐Releasing Hormone Neuronal Migration.” Seminars in Reproductive Medicine 25: 305–312. [DOI] [PubMed] [Google Scholar]
  37. Shu, M. , Hong D., Lin H., et al. 2022. “Single‐Cell Chromatin Accessibility Identifies Enhancer Networks Driving Gene Expression During Spinal Cord Development in Mouse.” Developmental Cell 57: 2761–2775. [DOI] [PubMed] [Google Scholar]
  38. Suter, T. , Blagburn S. V., Fisher S. E., Anderson‐Keightly H. M., D'Elia K. P., and Jaworski A.. 2020. “TAG‐1 Multifunctionality Coordinates Neuronal Migration, Axon Guidance, and Fasciculation.” Cell Reports 30: 1164–1177. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Suter, T. , and Jaworski A.. 2019. “Cell Migration and Axon Guidance at the Border Between Central and Peripheral Nervous System.” Science 365, no. 6456: eaaw8231. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Tang, W. , Ehrlich I., Wolff S. B., et al. 2009. “Faithful Expression of Multiple Proteins via 2A‐Peptide Self‐Processing: A Versatile and Reliable Method for Manipulating Brain Circuits.” Journal of Neuroscience 29: 8621–8629. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Taroc, E. Z. M. , Katreddi R. R., and Forni P. E.. 2020. “Identifying Isl1 Genetic Lineage in the Developing Olfactory System and in GnRH‐1 Neurons.” Frontiers in Physiology 11: 601923. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Taroc, E. Z. M. , Lin J. M., Tulloch A. J., Jaworski A., and Forni P. E.. 2019. “GnRH‐1 Neural Migration From the Nose to the Brain Is Independent From Slit2, Robo3 and NELL2 Signaling.” Frontiers in Cellular Neuroscience 13: 70. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Taroc, E. Z. M. , Naik A. S., Lin J. M., et al. 2020. “Gli3 Regulates Vomeronasal Neurogenesis, Olfactory Ensheathing Cell Formation, and GnRH‐1 Neuronal Migration.” Journal of Neuroscience 40: 311–326. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Taroc, E. Z. M. , Prasad A., Lin J. M., and Forni P. E.. 2017. “The Terminal Nerve Plays a Prominent Role in GnRH‐1 Neuronal Migration Independent From Proper Olfactory and Vomeronasal Connections to the Olfactory Bulbs.” Biology Open 6: 1552–1568. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Tessier‐Lavigne, M. , Placzek M., Lumsden A. G., Dodd J., and Jessell T. M.. 1988. “Chemotropic Guidance of Developing Axons in the Mammalian Central Nervous System.” Nature 336: 775–778. [DOI] [PubMed] [Google Scholar]
  46. Ware, M. , Hamdi‐Roze H., Le Friec J., David V., and Dupe V.. 2016. “Regulation of Downstream Neuronal Genes by Proneural Transcription Factors During Initial Neurogenesis in the Vertebrate Brain.” Neural Development 11: 22. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Wolfer, D. P. , Henehan‐Beatty A., Stoeckli E. T., Sonderegger P., and Lipp H. P.. 1994. “Distribution of TAG‐1/Axonin‐1 in Fibre Tracts and Migratory Streams of the Developing Mouse Nervous System.” Journal of Comparative Neurology 345: 1–32. [DOI] [PubMed] [Google Scholar]
  48. Wolman, M. A. , Sittaramane V. K., Essner J. J., Yost H. J., Chandrasekhar A., and Halloran M. C.. 2008. “Transient Axonal Glycoprotein‐1 (TAG‐1) and Laminin‐alpha1 Regulate Dynamic Growth Cone Behaviors and Initial Axon Direction In Vivo.” Neural Development 3: 6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Wray, S. 2001. “Development of Luteinizing Hormone Releasing Hormone Neurones.” Journal of Neuroendocrinology 13: 3–11. [DOI] [PubMed] [Google Scholar]
  50. Wray, S. , and Hoffman G.. 1986. “A Developmental Study of the Quantitative Distribution of LHRH Neurons Within the Central Nervous System of Postnatal Male and Female Rats.” Journal of Comparative Neurology 252: 522–531. [DOI] [PubMed] [Google Scholar]
  51. Wray, S. , Key S., Qualls R., and Fueshko S. M.. 1994. “A Subset of Peripherin Positive Olfactory Axons Delineates the Luteinizing Hormone Releasing Hormone Neuronal Migratory Pathway in Developing Mouse.” Developmental Biology 166: 349–354. [DOI] [PubMed] [Google Scholar]
  52. Wray, S. , Nieburgs A., and Elkabes S.. 1989. “Spatiotemporal Cell Expression of Luteinizing Hormone‐Releasing Hormone in the Prenatal Mouse: Evidence for an Embryonic Origin in the Olfactory Placode.” Brain Research. Developmental Brain Research 46: 309–318. [DOI] [PubMed] [Google Scholar]
  53. Yamamoto, M. , Boyer A. M., Crandall J. E., Edwards M., and Tanaka H.. 1986. “Distribution of Stage‐Specific Neurite‐Associated Proteins in the Developing Murine Nervous System Recognized by a Monoclonal Antibody.” Journal of Neuroscience 6: 3576–3594. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Yamamoto, M. , and Schwarting G.. 1991. “The Formation of Axonal Pathways in Developing Cranial Nerves.” Neuroscience Research 11: 229–260. [DOI] [PubMed] [Google Scholar]
  55. Yoshida, K. , Tobet S. A., Crandall J. E., Jimenez T. P., and Schwarting G. A.. 1995. “The Migration of Luteinizing Hormone‐Releasing Hormone Neurons in the Developing Rat Is Associated With a Transient, Caudal Projection of the Vomeronasal Nerve.” Journal of Neuroscience 15: 7769–7777. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Zouaghi, Y. , Alpern D., Gardeux V., et al. 2025. “Transcriptomic Profiling of Murine GnRH Neurons Reveals Developmental Trajectories Linked to Human Reproduction and Infertility.” Theranostics 15: 3673–3692. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Figure S1. Without tamoxifen injection Cntn2CreERT2 +/− have no spontaneous recombination. (a‐b) Cartoons representing the two experimental conditions, tamoxifen‐injected and non‐injected. (c‐d″) Immunofluorescent staining on injected or non‐injected E12.5 mice stained for the pan neuronal marker HuCD (green) and anti‐tdTomato antibody (magenta). HuCD‐labeled neurons in the nasal area included cells of the vomeronasal organ (VNO), olfactory epithelium (OE), and forebrain (FB). In the non‐injected animals (c‐c″), no tdTomato was detected in the neurons (empty arrowheads), while in tamoxifen‐injected animals (d‐d″), Cntn2Cre tracing (magenta) labeled neurons in the brain and nasal epithelia (white arrowheads). (g‐h″) Neurons of the trigeminal (TG) stained for HuCD (green) and Cntn2 tracing (magenta), consistently showing no Cntn2 tracing in non‐injected embryos (empty arrowheads), while extensive recombination was found in the tamoxifen‐treated embryos. Scale bars in f‐f′, 25 μm; g‐h″, 50 μm; c‐e″, 100 μm.

DVG-63-e70021-s002.tif (2.5MB, tif)

Appendix S1. Supplementary Information.

DVG-63-e70021-s001.xlsx (46.2KB, xlsx)

Data Availability Statement

The data that support the findings of this study are openly available in GEO at https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE234871, reference number GSE234871.

Articles from Genesis (New York, N.y. : 2000) are provided here courtesy of Wiley

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