Since RET can be activated by GFLs/GFRas either in cis or in trans (mainly by GDNF/GFRa1) in vitro, we asked if the expression patterns of Gfra1, Gfra2, Gdnf, and Nrtn in the developing SC and DRGs would provide insight into RET signaling in RA mechanoreceptors in vivo. We performed in situ hybridization for Ret, Gfra1, Gfra2, Gdnf, and Nrtn on embryonic day 13.5 (E13.5) and E15.5 wild-type DRG and SC sections. Double in situ hybridizations that characterize the expression of Gfra1 and Gfra2 in different populations of DRG neurons have been previously conducted (summarized in Figure 1—figure supplement 1K [Luo et al., 2009]).

Similar to previous characterization (Molliver et al., 1997; Luo et al., 2007, 2009), Ret is expressed in motor neurons and a mix of small and large diameter DRG neurons at E13.5 and E15.5 (Figure 1—figure supplement 1A–B). Most large diameter RET⁺ DRG neurons at these stages are the early RET⁺ DRG neurons, which develop into RA mechanoreceptors (Bourane et al., 2009; Luo et al., 2009). Gfra1 is highly expressed in some DRG neurons and motor neurons as well, but these GFRa1⁺ DRG neurons come from NTRK1⁺ precursors and are not early RET⁺ RA mechanoreceptors (Yu et al., 1998; Luo et al., 2009; Honma et al., 2010). In addition, Gfra1 is highly expressed in the dorsal root entry zone and the dSC, which are the target fields of the central projections of RA mechanoreceptors (Figure 1—figure supplement 1C–D). Gfra2 is expressed in a small number of large diameter DRG neurons, which were previously shown to be RA mechanoreceptors (Bourane et al., 2009; Luo et al., 2009), and some SC cells and motor neurons at these stages (Figure 1—figure supplement 1E–F; Oppenheim et al., 2000).

Nrtn is diffusely expressed at a low level in the SC and DRGs at both E13.5 and E15.5; Gdnf transcript is barely detected at E13.5 but is clearly expressed in DRG and motor neurons at E15.5 (Figure 1—figure supplement 1G–J). Thus, based on the expression patterns of RET signaling components in the developing SC and DRGs, RET in the central projections and cell bodies of developing RA mechanoreceptors could potentially be activated in cis by NRTN/ GFRa2 or in trans by GDNF/ GFRa1, which may come from neighboring DRG neurons, dorsal root entry zone cells, or dSC cells.

RA mechanoreceptors depend on RET for the growth of their third order central projections innervating layers III–V of SC. In postnatal Ret mutant mice, VGLUT1⁺ puncta, which label pre-synaptic terminals of mechanoreceptors and proprioceptors (Hughes et al., 2004; Paixão et al., 2013), are greatly reduced in layers III–V, indicating deficits in the third order central projections of RA mechanoreceptors (Luo et al., 2009). Since RA mechanoreceptors express a high level of Gfra2 but not any other Gfras (Luo et al., 2009), it is likely that RET in RA mechanoreceptors is activated by NRTN/GFRa2 in cis. Indeed, we previously found that Pacinian corpuscles, a subtype of RA mechanosensory end organs in the periphery, are not formed in Ret, Gfra2, or Nrtn mutant mice, supporting that NRTN/GFRa2-RET cis signaling occurs in RA mechanoreceptors (Luo et al., 2009). Here, we asked if NRTN-GFRa2/RET cis signaling is required for the growth of RA mechanosensory central projections as well. We performed immunostaining of VGLUT1 with postnatal day 7 (P7) Gfra2^GFP/GFP null and Nrtn^−/− null SC sections. No significant decrease of VGLUT1⁺ puncta in layers III–V of SC is observed in Gfra2 and Nrtn null mice (Figure 1A–C, Figure 1—source data 1 [p = 0.96], and data not shown). This result suggests that unlike RET signaling in the peripheral branches of RA mechanoreceptors, cis activation of RET by GFRa2 and NRTN may be dispensable for the normal development of central projections of RA mechanoreceptors.

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VGLUT1 dSC staining and RA mechanoreceptor number in P7 Gfra2 mutants.

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RA mechanoreceptor central projections and cell number in E13.5 Ret, Gfra1, Gfra2, and Nrtn mutants.

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To determine whether RA mechanoreceptors survive without Gfra2, we quantified the number of GFP⁺;NF200⁺ neurons per DRG section in P7 Gfra2^GFP/+ controls and Gfra2^GFP/GFP nulls. Green fluorescent protein (GFP) is expressed from the Gfra2 locus and most of GFP⁺;NF200⁺ neurons indicate RA mechanoreceptors in Gfra2^GFP mice. In agreement with our previous findings at P0 (Luo et al., 2009), we found a slight but non-significant decrease in RA mechanoreceptor number between controls and mutants (Figure 1D, Figure 1—source data 1 [p = 0.34]). Therefore, cis RET signaling via GFRa2 does not seem to be critical for the early postnatal survival of RA mechanoreceptors.

To understand the mechanism of RET signaling that controls growth of RA mechanosensory central projections, we genetically traced RA mechanoreceptors in Ret, Gfra1, Gfra2, and Nrtn mutant mice at different developmental stages. We first used Ret mutant mice, which serve as a positive control for the central projection deficit, to determine a robust method for visualizing RA mechanosensory interstitial branches at E13.5. We compared two methods that have been previously used. One is immunostaining of neurofilament-200 (NF200), which is expressed by large diameter DRG neurons, including RA mechanoreceptors, SA mechanoreceptors, and proprioceptors (Bourane et al., 2009). The other is to use a knockin/null allele of Ret (Honma et al., 2010), Ret^CFP (Uesaka et al., 2008), in which cyan fluorescent protein (CFP, a variant of GFP) is expressed from the Ret locus. Although Ret is expressed in both RA mechanoreceptors and some other DRG neurons at E13.5 (Luo et al., 2009), central projections of non-RA mechanoreceptor RET⁺ neurons, most of which develop into nociceptors, do not innervate the dSC until E15.5 or later (Ozaki and Snider, 1997). In addition, the expression of Ret in dSC neurons is not obvious until E15.5 (Figure 1—figure supplement 1B). Thus, the Ret^CFP allele may allow us to specifically visualize central projections of RA mechanoreceptors at E13.5.

To compare these two methods, we performed anti-NF200 and anti-GFP immunostaining on SC sections of E13.5 Ret^CFP/+ control and Ret^CFP/CFP null embryos. We observed a decrease in the density of NF200⁺ fibers in the dorsal horn (Figure 1—figure supplement 2B,E). This decrease of NF200⁺ central projections, however, is not dramatic. This is because the NF200 antibody also recognizes central projections of SA mechanoreceptors and proprioceptors, which develop in a manner temporally comparable to RA mechanoreceptors. In contrast, CFP⁺ fibers innervating the dSC display a dramatic reduction in Ret null mice (Figure 1—figure supplement 2C,F). Ret null CFP⁺ fibers reach the dorsal surface of the SC but rarely grow interstitial branches innervating layers III–V. We quantified the number of CFP⁺ pixels in the dorsal horn (displayed as percentage of CFP⁺ pixels normalized to the control) as a proxy for the extent of axon growth and found a significant decrease in CFP⁺ fibers in Ret mutant dorsal horn (Figure 1—figure supplement 2H and Figure 1—source data 2, [p < 0.001]). This result suggests that the Ret^CFP allele is a valid tool for visualizing central projection deficits of RA mechanoreceptors at E13.5.

Since Ret signaling can positively regulate the expression of its own signaling components or control neuronal survival (Luo et al., 2007; Baudet et al., 2008; Golden et al., 2010), it is conceivable that the lack of dSC CFP⁺ fibers could be due to a downregulation of CFP expressed in Ret null RA mechanoreceptors or death of RA mechanoreceptors. To exclude these possibilities, we quantified the number of CFP⁺ neurons in DRGs. We found that the number of CFP⁺ neurons per DRG section was not statistically different between Ret heterozygotes and null mice (Figure 1—figure supplement 2I and Figure 1—source data 2). In addition, the intensity of GFP⁺ fibers at the dorsal surface of the SC is comparable between Ret mutant and control mice. Therefore, the loss of CFP⁺ fibers in the dorsal horn of E13.5 Ret mutants must mainly be due to a deficit in growth of interstitial central axons, but not due to the down-regulation of CFP expression or the death of RA mechanoreceptors.

The finding that dSC VGLUT1 staining is largely normal in postnatal Gfra2 and Nrtn null mice suggests that cis RET signaling may be dispensable for RA mechanosensory central projections. To determine if the development of RA mechanosensory central projections depends on the trans activation of RET via GFRa1 and GDNF, we generated Gfra1 null (Gfra1⁻) mice (Figure 1—figure supplement 3A–B and ‘Materials and methods’). In situ hybridization of Gfra1 control and null DRG sections showed that Gfra1 transcripts are not produced in mice homozygous for this mutant allele (Figure 1—figure supplement 3C–D). In addition, kidneys are not formed in these Gfra1 null mice (data not shown), a phenotype consistent with previously reported Gfra1 null mice (Cacalano et al., 1998; Enomoto et al., 1998). Thus, the Gfra1⁻ allele we generated is a null allele.

If trans activation of RET via GFRa1 is required for the growth of interstitial central projections of RA mechanoreceptors, we expect to see a decrease of central projections of RA mechanoreceptors in the dSC of Gfra1 null mice. To test this idea, we generated E13.5 Gfra1^+/−;Ret^CFP/+ control and Gfra1^−/−;Ret^CFP/+ mutant embryos to examine RA mechanosensory central projections at this stage (Figure 1E–F). We found that innervation of dSC by CFP⁺ fibers was not reduced upon Gfra1 ablation (Figure 1G, Figure 1—source data 2). Additionally, the lack of Gfra1 function did not lead to a decrease of CFP⁺ DRG neurons (Figure 1H, Figure 1—source data 2). Together, our results suggest that trans activation of RET via GFRa1 is not required for the survival or central projection growth of RA mechanosensory neurons at E13.5.

Since no deficit was observed in the central projections of RA mechanoreceptors in E13.5 Gfra1 mutants, we next asked whether cis RET signaling is required for the initial growth of RA mechanosensory third order central projections. We crossed the Ret^CFP allele into Gfra2 and Nrtn null mice and examined central projections of RA mechanoreceptors at E13.5 (Figure 2A–D). In contrast to what we observed at P7, at this early development stage CFP⁺ central projections of RA mechanoreceptors are greatly reduced in both Gfra2 and Nrtn null SC sections (Figure 2E–F, Figure 1—source data 2, Gfra2 mutant has 9.50 ± 1.44% of control staining at thoracic levels [p < 0.001]). In addition, similar to the E13.5 Ret mutant mice, the number of CFP⁺ DRG neurons in Gfra2 and Nrtn null mice is comparable to that of control mice (Figure 2G–H, Figure 1—source data 2), suggesting that the loss of CFP⁺ fibers in the dSC of these mutant mice is due to a deficit in the interstitial central projection growth of RA mechanoreceptors. Thus, at E13.5, Gfra2 and Nrtn null mice phenocopy the central projection deficit of Ret mutant mice, which suggests that RET is activated by NRTN/GFRa2 in cis for the initial growth of RA mechanosensory central projections.

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If Ret, Gfra2, and Nrtn null mice phenocopy each other at E13.5, why do their postnatal VGLUT1 staining patterns look so different (Figure 1 and [Luo et al., 2009])? One possibility is that since Ret has a much broader expression pattern than Gfra2 in the dSC and DRGs, the dramatic loss of VGLUT1 staining in layers III–V of SC may be caused by the loss of RET signaling both in RA mechanoreceptors and other RET⁺ cells. For Gfra2 and Nrtn mutant mice, though central projection deficits of RA mechanoreceptors may persist postnatally, VGLUT1⁺ puncta from SA mechanoreceptors could mask the phenotype. Alternatively, RA mechanosensory central projections in Gfra2 and Nrtn mutant mice could recover at later developmental stages due to the function of other RET signaling mechanisms.

To differentiate these possibilities, we examined central projections of RA mechanoreceptors in Gfra2 null mice through development. We focused on Gfra2 instead of Nrtn mutant mice because: (1) the cell autonomous requirement of a co-receptor is the key to differentiate cis vs trans RET signaling; and (2) Gfra2 and Nrtn null mice display very similar phenotypes of RA mechanoreceptors. Since Ret begins to be expressed in additional populations of DRG neurons (Molliver et al., 1997; Luo et al., 2007) and dSC cells (Figure 1—figure supplement 1B) from E15.5, we can't use the Ret^CFP allele to visualize the central projections of RA mechanoreceptors at late developmental stages. To overcome this problem, we used a tandem allele (see ‘Materials and methods’ and Figure 3—figure supplement 1) of an inducible Cre allele of Ret (Ret^CreERT) and Rosa26 conditional red fluorescent protein (Rosa^Tdt). We combined these alleles with early (E11.5 and E12.5) 4-hydroxy tamoxifen (4-HT) treatment to specifically trace RA mechanoreceptors, as previously established (Luo et al., 2009).

We generated Gfra2^GFP/+; Ret^CreERT; Rosa^Tdt control and Gfra2^GFP/GFP; Ret^CreERT; Rosa^Tdt mutant mice and examined their SC and DRG sections at E15.5. Tdt⁺ fibers innervate layers III–V of the SC, which is consistent with specific genetic tracing of RA mechanoreceptors (Luo et al., 2009). In addition, the majority of Tdt⁺ DRG neurons are RET⁺, GFRa2⁺ (reported by the expression of GFP), but NTRK1⁻ at E15.5 (Figure 3A–J), further supporting the specific labeling of RA mechanoreceptors. We found that central projections of Gfra2 null RA mechanoreceptors are also decreased at E15.5 (Figure 3K–N, Figure 3—source data 1, Gfra2 mutant has 55.13 ± 2.82% of control staining at the thoracic level [p < 0.001]). Since the number of labeled DRG neurons is not significantly reduced in the mutant mice (Gfra2 mutants have 79.52 ± 8.39% of control cell number [p = 0.06]), the central projection phenotype mostly reflects a growth deficit at this developmental stage. Noticeably, the relative reduction of innervation in Gfra2 null mice at E15.5 is less severe compared to that of E13.5 mutants (Figure 2), suggesting that central projections of Gfra2 null RA mechanoreceptors may start to recover at this stage.

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RA mechanoreceptor central projections and cell number in E15.5 Gfra2mutants.

https://doi.org/10.7554/eLife.06828.011

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To determine if RA mechanoreceptors require Ret but not Gfra2 for their central projection growth at later developmental stages, we generated E18.5 Ret^CreERT/+;Rosa^Tdt control and Ret^{CreERT/CreERT};Rosa^Tdt mutant embryos (Ret^CreERT is a null allele of Ret). Consistent with previous results (Bourane et al., 2009; Luo et al., 2009; Honma et al., 2010), we found that RA mechanosensory central projections are greatly reduced in the Ret mutant mice (Figure 4A,C,I, Figure 4—source data 1, Ret mutant has 35.86 ± 4.97% of control staining at thoracic levels [p < 0.001]). In addition, we counted the number of Tdt⁺ neurons in L4/L5 DRGs and found that the number of Tdt⁺ RA mechanoreceptors is dramatically reduced as well (Figure 4B,D,J, Ret mutant has 52.52 ± 7.76% of control cell number [p < 0.001]). Taken together, these results suggest that Ret is absolutely required for both survival and central projection growth of RA mechanoreceptors at E18.5.

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RA mechanoreceptor central projections and cell number in E18.5 Ret, Gfra2, Gfra1, and Gfra1;Gfra2 mutants.

https://doi.org/10.7554/eLife.06828.014

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In contrast, central projections of Tdt⁺ Gfra2 null RA mechanoreceptors are only slightly reduced at E18.5 (Figure 4E,G,I, Figure 4—source data 1, Gfra2 mutant has 86.34 ± 4.48% of control staining at thoracic levels [p = 0.01]). At P7, almost no difference is observed (data not shown). Similarly, the number of Tdt⁺ RA mechanoreceptors is only slightly reduced in Gfra2 null mice (Figure 4F,H,J Gfra2 mutant has 84.01 ± 5.16% of control cell number [p = 0.04]), indicating that extensive cell death of RA mechanoreceptors resulting from an absence of RET signaling does not occur in Gfra2 null mice. The discrepancy between E18.5 Ret and Gfra2 mutant phenotypes suggests that RET signaling still occurs in neonatal Gfra2 null RA mechanoreceptors. To demonstrate this, we quantified the expression of phospho-S6 ribosomal protein, which is downstream of RET/PI3K/mTOR signaling (Plaza-Menacho et al., 2010), in RA mechanoreceptors. We found that the proportion of GFP⁺ RA mechanoreceptors which express phospho-S6 in P0 Gfra2^GFP/+ control and Gfra2^GFP/GFP mutant DRGs was similar (Figure 4—figure supplement 1 [p = 0.51]). This result is consistent with the idea that RET activation occurs in neonatal RA mechanoreceptors without Gfra2.

Collectively, our results suggest that Gfra2 null RA mechanoreceptors display a central projection deficit at E13.5 but recover during later development, which explains the almost normal VGLUT1 staining in layers III–V of SC at P7. In addition, our data indicate that from E15.5, an additional GFRa2 independent but RET-dependent mechanism begins to play a role in promoting the survival and central projection growth of RA mechanoreceptors.

To determine if this GFRa2-independent but RET-dependent mechanism requires GFRa1, we examined genetically labeled Gfra1^+/−;Ret^CreERT/+;Rosa^Tdt control and Gfra1^−/−;Ret^CreERT/+;Rosa^Tdt mutant SC and DRGs at E18.5. Similar to E13.5, neither RA mechanosensory central projections nor their number is significantly decreased in Gfra1 null mice (Figure 5A–D,I–J, Figure 4—source data 1), suggesting that simply disrupting trans activation of RET via GFRa1 is not sufficient to block Ret signaling in RA mechanoreceptors.

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The lack of Ret-mutant-like survival and central projection phenotypes of RA mechanoreceptors in both Gfra1 and Gfra2 single null mice made us wonder if cis and trans RET signaling function in the same developmental process and thus loss of one co-receptor can be compensated for by the other. To test this idea, we generated Gfra1;Gfra2 double knockout mice, in which RA mechanoreceptors were specifically labeled with Tdt using the Ret^CreERT;Rosa^Tdt tandem allele. We examined control and double null SC sections and DRGs at E18.5. We found that Tdt⁺ RA mechanosensory central projections are greatly reduced in the dSC (Figure 5E,G,I, Figure 4—source data 1, Gfra1;Gfra2 double mutant has 27.25 ± 2.09% of control staining at thoracic levels [p < 0.001]). In addition, fewer Tdt⁺ RA mechanoreceptors remain in the double knockout DRGs (Figure 5F,H,J, Gfra1;Gfra2 double mutant has 38.17 ± 2.65% of control cell number [p < 0.001]), indicating that a significant number of RA mechanoreceptors die in the absence of Gfra1 and Gfra2. Strikingly, the extent of reduction in both cell number and central projections of RA mechanoreceptors is comparable between the Ret null and Gfra1:Gfra2 double null mice. Thus, our in vivo analyses strongly suggest that RET in RA mechanoreceptors is activated via both GFRa1 and GFRa2.

Is RET in RA mechanoreceptors activated by GFRa1 in cis or trans? Although Gfra1 is not widely expressed in RA mechanoreceptors in wild-type mice, could it be upregulated to compensate for the loss of Gfra2 in the Gfra2 null mice? To address these questions, we conducted double fluorescent in situ hybridization of Gfra1 and GFP with E14.5 Gfra2^GFP/+ control and Gfra2^GFP/GFP null DRG sections. We found that a comparable low number of GFP⁺ neurons expressed Gfra1 transcript in both mutants and controls (Figure 6A–C [p = 0.52]), suggesting that Gfra1 is not upregulated in Gfra2 null RA mechanoreceptors. In addition, we performed in situ hybridization of Gfra1 with P0 Gfra2^GFP/+;Ntrk1^+/− control, Gfra2^GFP/+;Ntrk1^−/− null, and Gfra2^GFP/GFP;Ntkr1^−/− double null DRG sections. We previously showed (Luo et al., 2009) that Gfra1 is expressed in NTRK1⁺ DRG neurons and that the expression of Gfra1 is completely lost in Ntrk1 null mice. Here, we found that while Gfra1 expression was observed in Gfra2^GFP/+;Ntrk1^+/− control DRGs, no Gfra1 expression was observed in either Gfra2^GFP/+;Ntkr1^−/− null or Gfra2^GFP/GFP;Ntkr1^−/− double null DRGs (Figure 6D–F). This result indicates that the expression of Gfra1 in Gfra2 null DRG neurons still fully depends on NTRK1 signaling and thus it must be expressed in the non-RA mechanoreceptors. Moreover, we performed quantitative RT-PCR (QPCR) for Gfra1 transcripts in DRGs from E13.5, E15.5, and E18.5 Gfra2^GFP/+ control and Gfra2^GFP/GFP mutant embryos. We found no significant difference in the expression of Gfra1 between control and mutant DRGs at any stage (Figure 6—figure supplement 1, Figure 6—source data 1), suggesting that Gfra1 is not transcriptionally upregulated in DRG neurons upon Gfra2 ablation.

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QPCR of Gfra1 in embryonic Gfra2 null DRGs.

https://doi.org/10.7554/eLife.06828.018

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Although Gfra1 transcript in most RA mechanoreceptors is below the detection level of in situ hybridization, it remains possible that an undetectable amount of GFRa1 could function in cis to promote RET signaling in RA mechanoreceptors. To exclude this possibility and to demonstrate that RET in RA mechanoreceptors is indeed activated by GFRa1 in trans, we used DRG explants from E14.5 embryos of different mutant backgrounds and treated these explants with NRTN, GDNF, GFRa1 plus GDNF, or GFRa1 alone.

In E14.5 explants harboring the Ret^CFP allele, the cell bodies and axons of RET⁺ neurons, some of which are RA mechanoreceptors, can be identified by anti-GFP immunostaining. We found that CFP⁺ neurons in Ret^CFP/+ control DRG explants grow long axons upon NRTN, GDNF, or GFRa1 plus GDNF, but not GFRa1 alone treatment (Figure 7—figure supplement 1A–D,I, and Figure 7—source data 1). In addition, the number of CFP⁺ DRG neurons is reduced in GFRa1 alone culture (Figure 7—figure supplement 1J, Figure 7—source data 2), suggesting that either cell death or down-regulation of Ret, and thus CFP expression, occur in the absence of RET signaling. Similarly, CFP⁺ neurons in Ret^CFP/CFP null DRG explants lost their responsiveness to GFLs completely (Figure 7—figure supplement 1E–H,I–J, Figure 7—source data 2), suggesting that this assay reflects RET-dependent signaling.

Next, we examined DRG explants harboring the Gfra2^GFP allele, which drives a much lower level of GFP expression than Ret^CFP. Although some small diameter DRG neurons also express Gfra2 around P0 or later (Luo et al., 2007), in this Gfra2^GFP mouse line GFP is mainly detected in the large diameter RA mechanoreceptors (Luo et al., 2009), which express a much higher level of Gfra2. Therefore, anti-GFP staining of E14.5 Gfra2^GFP DRG explants should specifically show RA mechanoreceptors. Since GFP⁺ axons of these explants could not be reliably imaged and quantified due to the low level of GFP expression, we approximated the extent of RET signaling in Gfra2^GFP DRG explants by quantifying the number of discernable GFP⁺ cell bodies. We found that Gfra2^GFP/+ control DRG neurons show robust responses upon GFL application (Figure 7A–D,Q, Figure 7—source data 2). Interestingly, Gfra2^GFP/GFP null DRG neurons lost their responsiveness to NRTN, but retain GFP expression in the presence of either GDNF or GFRa1 plus GDNF (Figure 7E–H,Q, Figure 7—source data 2). These results suggest that a GFRa2-independent but RET-dependent mechanism can mediate GDNF responsiveness of RA mechanoreceptors.

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Quantification of axonal growth in Ret mutant DRG explants.

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GFP⁺ neuron number in Gfra2 null and Gfra2;Ntrk1 double null explants.

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Densimetric measurements of GFRa1 in DRG cell extracts and supernatants.

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How can Gfra2 null RA mechanoreceptors retain their responsiveness to GDNF? It could be due to: (1) a very low level of GFRa1 is expressed in RA mechanoreceptors, which activates RET in cis in the presence of GDNF; or (2) GFRa1 expressed by neighboring DRG neurons binds GDNF and activates RET in RA mechanoreceptors in trans. To differentiate between these possibilities, we cultured E14.5 Gfra2^GFP/GFP;Ntrk1^−/− double mutant DRGs. Since the expression of Gfra1 in non-RA mechanoreceptor DRG neurons fully depends on NTRK1 signaling, as shown previously (Luo et al., 2009) and above (Figure 6D–F), GFRa1 should be depleted from non-RA mechanoreceptors in Gfra2^GFP/GFP;Ntrk1^−/− double mutant DRGs. Therefore, if GFRa1 is expressed at a low level in RA mechanoreceptors and activates RET in cis, the double null explants should retain their responsiveness to GDNF. On the other hand, if GFRa1 expressed by neighboring neurons activates RET in RA mechanoreceptors in trans, the GDNF responsiveness would be lost in the Gfra2;Ntrk1 double nulls. Here, we found that Gfra2^GFP/+;Ntrk1^−/− control explants were responsive to NRTN, GDNF, and GDNF plus GFRa1, but not GFRa1 alone (Figure 7I–L,R, Figure 7—source data 2). In contrast, Gfra2^GFP/GFP;Ntrk1^−/− double null explants only respond to GDNF plus GFRa1, but not to NRTN, GDNF, or GFRa1 (Figure 7M–P,R, Figure 7—source data 2). The loss of responsiveness of RA mechanoreceptors to GDNF in Gfra2;Ntrk1 double null DRG explants strongly suggests that Gfra1 is not expressed at a functional level in RA mechanoreceptors and that RET in Gfra2 null RA mechanoreceptors is activated by exogenous GFRa1 from the neighboring DRG neurons in trans.

Trans activation of RET could occur by direct contact between membranes of cells which express either RET or GFRa1, or by soluble GFRa1 which is shed from the cell surface. To determine whether GFRa1 is released by DRG neurons, we cultured dissociated DRGs from E18.5-P1 wild-type, Gfra2^−/−, and Gfra1^−/− mice. We collected cell lysates and concentrated media from days 3–6 in vitro and then performed Western blot analysis.

Immunoblotting with anti-GFRa1 revealed a doublet at ∼55–65 kDa in wild-type and Gfra2^−/− cell lysates, which was absent in the Gfra1^−/− samples (Figure 7S, lanes 1–3), confirming the specificity of the anti-GFRa1 antibody. A positive band of ∼55 kDa was present in concentrated supernatants of wild-type and Gfra2^−/− but not Gfra1^−/− cultures (Figure 7S, lanes 4–6), suggesting that soluble GFRa1 is shed from neonatal DRG cells. Together with reports of GFRa1 being shed by Sciatic nerve Schwann cells, immortalized neuronal progenitors (Paratcha et al., 2001), and adult DRG explants (He et al., 2014), our findings indicate that GFRa1 can be released by many cell types during both developmental and adult stages. Therefore, it is possible that RET in RA mechanoreceptors is activated in trans by both soluble GFRa1 and GFRa1 tethered to the membranes of neighboring cells.

In addition, although there is no significant increase of Gfra1 transcripts in Gfra2 null DRGs by in situ or QPCR (Figure 6 and Figure 6—figure supplement 1), it remains possible that post-transcriptional regulation may occur to alter the translation, perdurance, or release of GFRa1. To test this possibility, we quantified the amount of GFRa1 in cell lysates and supernatants of wild-type and Gfra2^−/− cultures by densitometry. We found that the amount of GFRa1 expressed in the cell or shed into the media was not significantly different between wild-type and Gfra2 null cultures (Figure 7U–V, Figure 7—source data 3). Therefore, Gfra2 null DRGs do not produce or release more GFRa1 protein to compensate for the loss of Gfra2.

We also investigated whether GFRa2 is normally shed by DRGs. The specificity of the anti-GFRa2 antibody was confirmed by the absence of a ∼75 kDa band from Gfra2 null cell lysates, which was present in both wild type and Gfra1 null cultures (Figure 7T, lanes 1–3). Furthermore, secreted GFRa2 band was also present in the supernatants of wild-type and Gfra1 DRG cultures, but not in Gfra2 null cultures. Therefore, both GFRa1 and GFRa2 are normally released by DRGs during early postnatal development.

As described above, the central projections of Gfra2 null RA mechanoreceptors display a severe, Ret-like deficit at E13.5, but begin to recover from E15.5, which is due to compensation by trans signaling via GDNF/GFRa1. Why is trans RET signaling able to compensate for the loss of cis signaling during late embryonic development, but not at E13.5? One possible reason for the delay is the availability of trans signaling components. Our in situ hybridization data suggest that Gfra1 is expressed at high levels at both E13.5 and E15.5, but the expression of Gdnf is greatly increased in DRGs from E13.5 to E15.5 (Figure 1—figure supplement 1). To provide additional evidence for the dynamic expression of Gdnf during development, we examined DRG and SC sections of E13.5 and E15.5 Gdnf^LacZ/+ (Moore et al., 1996) embryos using X-Gal staining. We found that the expression of LacZ increased significantly in DRGs from E13.5 to E15.5 (Figure 8A–E [p < 0.001], Figure 8—figure supplement 1). In addition, X-Gal staining was found in the E15.5 dorsal root, the pathway through which DRG central projections travel to reach the dSC (Figure 8A–D, black arrows). Thus, the expression of Gdnf seems to significantly increase in both the DRG and dorsal root from E13.5 to E15.5, providing a possible explanation for why the trans compensation occurs from E15.5.

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Previous expression analyses revealed that Gfra1 is expressed more broadly than Ret, and cells which express Gfra1 usually lie adjacent to Ret-expressing cells (Trupp et al., 1997; Yu et al., 1998). This expression pattern suggests that GFRa1 may have RET-independent functions or that GFRa1 may interact with RET expressed on the surfaces of other cells in trans. Indeed, evidence for both ideas has been demonstrated. GFRa1 and GDNF interact with NCAM in neurons and Schwann cells to promote neurite outgrowth and Schwann cell migration (Paratcha et al., 2003; Nielsen et al., 2009). Additionally, GFRa1 and GDNF are required for the proper migration of cortical GABAergic interneurons and can act as ligand-dependent adhesion molecules for synapse formation, independent of RET and NCAM (Pozas and Ibáñez, 2005; Ledda et al., 2007). Recently, it was also shown that GFLs have additional roles in cortical development via interactions with Syndecan-3, likely independent of RET, GFRas, and NCAM (Bespalov et al., 2011). On the other hand, RET can be activated by GFRa1 and GDNF in trans using both heterologous cells and tissue explants (Paratcha et al., 2001; Ledda et al., 2002; Patel et al., 2012; He et al., 2014). Trans RET signaling may affect many cellular processes, including directional axonal outgrowth and promotion of axon regeneration (Paratcha et al., 2001; Airaksinen and Saarma, 2002; Ledda et al., 2002).

Evidence for physiologically relevant in vivo function of trans RET signaling, however, has remained less conclusive. Enomoto et al. (2004) generated a ‘cis-only’ mouse model in which Gfra1 is expressed in all RET-expressing cells, but not in cells that do not express RET. Using this model, they found that the major RET-dependent developmental processes were completely normal, suggesting that trans signaling is likely to be irrelevant for most RET-dependent processes. Results from this model, however, may not necessarily preclude a physiological role for trans signaling. Not only does this model present a loss of trans signaling, but it also presents a gain of function: Gfra1 is expressed at a high level in RET⁺ cells which may not normally express this co-receptor. If cis and trans RET activation lead to similar physiological outcomes, any deficits due to the loss of trans signaling may be masked by a gain of cis signaling. Indeed, the gain of function of GFRa1 was recently demonstrated in enteric hematopoietic cells derived from cis-only mice (Patel et al., 2012). Thus, whether trans RET signaling has any physiological function in development has remained an open question.

We aimed to address this question by analyzing the survival and growth of RA mechanosensory central projections using loss-of-function mouse lines. Here, we found that loss of cis signaling, via ablation of either Gfra2 or Nrtn, produces a central projection deficit during early embryonic development (Figure 2). Our findings are consistent with previous observations using a different Ret knock-in line and NRTN ectopic expression (Honma et al., 2010), but differ from the findings using anti-GFRa2 staining to visualize RA mechanosensory central projections at E13.5 (Bourane et al., 2009). This is likely due to different subcellular localization of CFP and GFRa2, as well as the expression of GFRa2 by some dSC cells (Figure 1—figure supplement 1E,F), which may mask the RA mechanoreceptor phenotype. Interestingly, this phenotype recovers during late embryonic development and the central projections of Gfra2 null mice seem nearly normal postnatally (Figures 1, 4). Thus, our results suggest that cis RET signaling is required for the initial growth of RA mechanosensory central projections, but an additional cis signaling independent process takes place during later development. Indeed, the loss of both cis and trans signaling in Gfra1;Gfra2 double mutants recapitulates the Ret phenotype (Figures 5, 6). Furthermore, using DRG explant and dissociated culture, we demonstrated that soluble GFRa1 is normally released by DRGs and that GFRa1 produced by NTRK1⁺ DRG neurons present a potential source to activate RET in RA mechanoreceptors in trans to promote their survival (Figure 7). Taken together, our results suggest that trans RET signaling contributes to the development of RA mechanoreceptors in vivo.

Nevertheless, the exact subcellular locus of trans RET activation in RA mechanoreceptors remains speculative. The expression pattern of Gfra1 suggests that trans RET activation is possible in the axons of RA mechanoreceptors along their path to dSC, and/or at the cell body within the DRG. Although individual DRG cell bodies are surrounded by satellite glial cells, large macromolecules and proteins are able to invade the space between the neuron and satellite cell (Hanani, 2005), suggesting trans RET activation by soluble GFRa1 could occur within DRGs.

Signaling of neurotrophic RTKs, such as NTRK1, NTRK2, and NTRK3, is critical for the specification and survival of numerous classes of neurons (Ernsberger, 2009). RET signaling also plays important roles in survival, differentiation, and specification of distinct neuronal classes (Enomoto, 2005). For example, RET signaling components are absolutely required for the survival of enteric neurons (Taraviras et al., 1999), but their roles in DRG neuron survival are more complicated to dissect. Previously, it was reported that the number of total DRG neurons is not significantly reduced in neonatal and early postnatal Ret mutants (Luo et al., 2007). At that time, specific molecular markers or genetic approaches for labeling RA mechanoreceptors had not been identified, so it was impossible to specifically assay the role of RET signaling in the survival of this neuronal population. Given that RA mechanoreceptors represent a very small proportion of the total DRG neurons (Molliver et al., 1997; Luo et al., 2007), a partial loss of this population may not lead to a significant change in cell counts of total DRG neurons.

In another paper (Luo et al., 2009), it was proposed that RA mechanoreceptors depend on NRTN-GFRa2/Ret signaling for their development. This was based on the findings that GFRa2 is the only co-receptor expressed in RA mechanoreceptors and that Ret, Gfra2, and Nrtn null mice display the same no-Pacinian-corpuscle phenotype. Since RET can't be used as a molecular marker to quantify the number of RA mechanoreceptors in Ret null mice, the number of P0 RET⁺/NTRK1⁻ and Gfra2^GFP DRG neurons, most of which indicate the RA mechanoreceptors, was quantified in Nrtn and Gfra2 nulls. No significant change in cell number between mutants and controls was found, suggesting that Gfra2 and Nrtn are not required for survival of neonatal RA mechanoreceptors. These results are interesting in light of the current findings. Here, we show that when cis signaling via NRTN/GFRa2 is perturbed (as was tested in [Luo et al., 2009]), trans signaling via GDNF/GFRa1 can activate RET in RA mechanoreceptors to support their survival and central projection growth. When the number of genetically labeled RA mechanoreceptors was quantified in different mutant backgrounds, we found only marginal changes in Gfra2 nulls but drastic decreases in Ret mutants at E18.5 (Figure 4). The slight difference between the current and previous findings regarding the loss of RA mechanoreceptors in Gfra2 nulls at P0 (Luo et al., 2009) is likely due to different methods in identifying and quantifying RA mechanoreceptors. In short, the current study clarifies that RET signaling is required for RA mechanoreceptor survival but simple disruption of cis RET signaling components may not reveal this deficit.

What is the purpose for RET to be activated both in cis and in trans? Do cis and trans signaling activate different cellular responses and influence distinct developmental processes, or do cis and trans signaling exert similar physiological effect? Here, we found that, in RA mechanoreceptors, cis and trans signaling seem to produce similar biological outputs in vivo. Our results demonstrate that cis and trans signaling can compensate for the loss of each other to promote both the central projection growth and survival of RA mechanoreceptors (Figure 8F). This compensatory ability suggests that the existence of both cis and trans activation is likely to enhance but not diversify outcomes of RET signaling. Consistent with this notion, a recent study found that peripheral nerves secrete both GDNF and GFRa1, which attracts perineural invasion of heterogeneous cancer cells, some of which expresses Ret and Gfras, while some express only Ret (He et al., 2014).

In an attempt to show that GFRa1 is normally released from wild-type DRG cells for trans RET signaling, we found the same for GFRa2 (Figure 7). Given that soluble GFRa2 could also activate RET in trans with NRTN (Worley et al., 2000), our finding raises many interesting questions, such as whether all GFRas are secreted and whether ‘cis’ and ‘trans’ RET signaling normally co-exist even when RET and GFRas are expressed in the same cells. It seems plausible that even for GFRas co-expressed with RET (usually defined as ‘cis’ signaling), such as GFRa2 in RA mechanoreceptors, it could be secreted and then upon NRTN binding activates RET in the cell from which it was released (‘trans’ activation).

Although cis and trans activation of RET lead to a similar biological outcome in the growth and survival of RA mechanoreceptors, it is worth noting that substantial differences likely exist between the signaling processes of cis and trans RET activation. Cis RET activation might be more efficient, given that GFRas and RET are located in the same membrane. In addition, cis and trans signaling could differ in the kinetics of recruitment of RET to lipid rafts upon GFLs stimulation, interactions with downstream-associated proteins, and the longevity of activated RET and downstream effectors (Tansey et al., 2000; Paratcha et al., 2001). Additionally, it remains possible that although the gross structure of the RA mechanosensory central projections seems mostly normal in mice lacking either cis or trans signaling, more precise aspects of circuit formation, such as specific synapse formation, which are beyond the resolution of current analysis, may differentially depend on cis or trans signaling. Finally, it will be interesting to see whether cis and trans signaling can produce similar biological outcomes in other systems, as we have shown here for RA mechanoreceptors. Future experiments to carefully dissect cis and trans RET signaling in other types of cells and tissues will address these issues.

Mice except GDNF^lacZ line were raised in a barrier facility in Hill Pavilion, the University of Pennsylvania. All procedures were conducted according to animal protocols approved by Institutional Animal Care and Use Committee (IACUC) of the University of Pennsylvania and National Institutes of Health guidelines. GDNF^lacZ mice were raised in accordance with the European Community Council Directive of November 24, 1986 (86/609/EEC), and approved by the ethics. Most mice used in this paper were described previously: Ret^CreERT, Ret^CFP, Nrtn^+/− (purchased from the Jackson lab), Gfra2^GFP (re-derived using sperm provided by Dr Jeffery Milbrandt at the Washington University), Rosa26^Tdt, and Gdnf^LacZ mice (Moore et al., 1996; Heuckeroth et al., 1999; McDonagh et al., 2007; Uesaka et al., 2008; Luo et al., 2009; Madisen et al., 2010). The Ntrk1⁻ allele was generated by crossing the floxed TrkA^F592A allele (Chen et al., 2005) to germline Cre mice. The generation of Gfra1 conditional and null mice and the Ret^CreERT;Rosa^Tdt tandem allele are described below. All mice except Gdnf^LacZ were maintained on a mixed C57BL/6J and CD1 background. Gdnf^LacZ mice were maintained on a C57BL/6N background. Except for Gfra1;Gfra2 double null animals (n = 2), at least three animals per genotype were examined. N-values for explants are listed in Figure 7—source data 1, 2.

We generated Gfra1 conditional knockout mice, in which loxP sites flank exon 6 of Gfra1, by homologous recombination. Mice harboring the floxed allele were crossed to germ line Cre mice, resulting in a Gfra1 allele lacking exon 6. The loss of Gfra1 transcript in Gfra1^−/− mice was confirmed by in situ hybridization of mutant and control DRGs (see Figure 1—figure supplement 3).

Since Ret and Rosa26 loci are located only ∼5 megabases apart on mouse chromosome 6, we generated a tandem configuration of Ret^CreERT and Rosa^Tdt alleles, which are linked during meiosis and became a great genetic advantage for our experiments (Figure 3—figure supplement 1). We used this tandem Ret^CreERT;Rosa^Tdt allele to specifically label RA mechanoreceptors in different mutant mouse lines described in the text.

We set up timed pregnancy mating for mice in the evening and checked mice for vaginal plugs the following morning. The time when a female mouse was found to have a plug was counted as embryonic day 0.5 (E0.5). We treated embryos harboring the Ret^CreERT;Rosa^Tdt reporter allele with 4-hydroxy-tamoxifen (4-HT, 2 mg and 1 mg at E11.5, and E12.5, respectively) by oral gavage to pregnant female mice to specifically label RA mechanoreceptor population.

Spinal columns of embryos and neonatal mice at the desired developmental stages were dissected out and directly immersed in PBS/4% paraformaldehyde (PFA) for 2 to 4 hr at 4°C. Postnatal mice were sacrificed with CO₂, transcardially perfused with 4% PFA, and spinal columns were dissected out and post-fixed with 4% PFA for 2 hr at 4°C. They were then cryo-protected in 1× PBS, 30% sucrose overnight. 20-µm frozen sections of SC and DRGs were cut using a CM1950 cryostat (Leica, Buffalo Grove, IL). Immunostaining of SCs and DRG sections were performed as described previously (Niu et al., 2013). Antibodies used are as follows: rabbit anti-GFP (A-11122, 1:2000, Invitrogen, Carlsbad, CA), chicken anti-GFP (GFP-1020, 1:1000, Aves, Tigard, OR), chicken anti-NF200 (NF-H, 1:500, Aves), rabbit anti-NF200 (N4142, 1:1000, Sigma, St. Louis, MO), rabbit anti-cRet (18121, 1:50, IBL, Minneapolis, MN), rabbit anti-NTRK1 (06-574, 1:1000, Millipore, Temecula, CA), guinea pig anti-VGLUT1 (AB5905, 1:1000, Millipore), rabbit anti-phospho-S6 (2215s, 1:200, Cell Signaling, Beverly, MA), and Alexa Fluorescent conjugated Goat or Donkey secondary antibodies (1:500, Invitrogen or Jackson ImmunoResearch, West Grove, PA).

Embryos of the desired age were eviscerated and fixed in 1% PFA, 2 mM MgCl₂, 5 mM EGTA, 0.02% NP40, and 0.2% glutaraldehyde in phosphate buffer (pH 7.4) for 1.5 to 2 hr at 4°C. Vibratome sections were incubated for 30 min in washing solution (2 mM MgCl₂, 0.02% NP-40 in phosphate buffer pH 7.4). LacZ reaction was developed with X-gal (1 mg/ml) at 37°C.

DIG- or FITC-labeled riboprobes were synthesized using a DIG or FITC RNA labeling kit (11175025910, Roche, Indianapolis, IN). Template for GFP was amplified by PCR and subcloned into vector pGEM-T Easy (A1360, Promega, Madison, WI). Antisense RNA probes for Ret, Gfra1, Gfra2, Gdnf, and Nrtn were generated as previously described (Luo et al., 2009). The detailed procedures of in situ hybridization and double fluorescent in situ hybridization were performed as described previously (Fleming et al., 2012).

DRGs from E13.5, E15.5, and E18.5 Gfra2^GFP/+ and Gfra2^GFP/GFP embryos were dissected and rapidly frozen on dry ice. RNA was extracted with the GeneJet RNA Purification Kit (K0731, Fermentas, Vilnius, Lithuania) and cDNAs were generated using Super-Script III First-Strand Synthesis System (18080-51, Invitrogen). 500 ng of total RNA was used for each RT reaction in a total volume of 25 μl. QPCR reactions were performed in triplicate for three samples of each age and genotype. QPCR reactions contained SYBR Green PCR master mix (4309155, Life Technologies, Carlsbad, CA), 0.5 μM of each primer, and 3 μl (for Gfra1) or 1 μl (for Gapdh) of cDNA template per 15 μl reaction. Reactions were run and analyzed on a StepOnePlus Real-Time PCR System (Applied Biosystems, Carlsbad, CA). Primers used were Gapdh (5′-CCACCAACTGCTTAGCCCCC-3′ and 5′-GCAGTGATGGCATGGACTGTGG-3′) and Gfra1 (5′-TGTCTTTCTGATAATGATTACGGA-3′ and 5′-CTACGATGTTTCTGCCAATGATA-3′). p-values between samples were calculated from ΔCT values with the Student's t-test, and relative concentrations were calculated by the 2^−ΔΔCT method (Livak and Schmittgen, 2001).

E14.5 embryos were removed from the dam and placed in F-12 media (11765-047, Invitrogen) on ice. SCs with attached DRGs were dissected from the spinal column, and individual DRGs were removed and placed in fresh F-12 on ice. Using a dissecting needle, DRGs were cleaned and bisected, and then placed in fresh F-12. Explants were grown on Superfrost Plus slides (22-034-979, Fisher, Waltham, MA) coated with poly-L-lysine (P1274, Sigma, 0.1 mg/ml in ddH2O overnight at 4°C) and laminin (354232, BD, Franklin Lakes, NJ), 20 μg/ml in HBSS [14170122, Invitrogen] at 37°C for one to 3 hr). Immediately before placing explants on the slide, slides were washed with HBSS and DRG culture medium (Neurobasal medium [21103-049, Invitrogen], 1× B27 [17504-044, Invitrogen], 100 U/ml penicillin/streptomycin [15140-122, Invitrogen], 2 mM L-glutamine [25030-081, Invitrogen], and 35 mM glucose). DRG culture media supplemented with the appropriate recombinant proteins (50 ng/ml Nrtn [477-MN-025, R&D, Minneapolis, MN], 100 ng/ml GDNF [512-GF-010, R&D], 300 ng/ml GFRa1 [560-GF-100, R&D], or 100 ng/ml GDNF and 300 ng/ml GFRa1) were added to the culture dish. Four to six DRG explants were placed on each slide and the culture dishes were carefully moved to a 37°C incubator and left undisturbed overnight. Following 16–24 hr of incubation, cultures were rinsed with PBS and fixed in 4% PFA in PBS for 30 min at room temperature. Immunofluorescence was then performed directly in the culture dish using antibody dilutions described above. Following secondary antibody, explant slides were mounted on microscope slides using Superglue, and coverslipped with Fluoromount-G (0100-01, Southern Biotech, Birmingham, AL) and 22 × 22 mm coverglass.

DRGs from E18.5-P1 mice were collected into HBSS on ice. DRGs were first digested in 0.5 mg/ml collagenase (LS4186, Worthington, Lakewood, NJ) plus 100 U/ml penicillin/streptomycin, 10 mM HEPES, and 1× MEM vitamins (M6895, Sigma) in MEM (11095072, Invitrogen) at 37°C for 30 min, followed by a second digestion with 0.05% trypsin (25200056, Invitrogen) plus 100 U/ml penicillin/streptomycin, 10 mM HEPES, and 1× MEM vitamins in MEM at 37°C for 30 min. Digestion was stopped by adding 5% FBS and 10 mM HEPES in HBSS. Cells were then triturated with a fire polished Pasteur pipette to a homogenous solution. The cells were then pelleted at 500×g for 5 min and resuspended in DRG culture media, as described above, supplemented with 50 ng/ml NRTN, 100 ng/ml GDNF, and 50 ng/ml NGF (556-NG-100, R&D). Cells were plated in six-well collagen-coated plates (Millipore, PICL06P05) and cultured at 37°C and 5% CO₂. After 2 days, media were removed and cells were rinsed with warmed Neurobasal media. 2 ml of fresh DRG culture media supplemented with NRTN, GDNF, and NGF (but without B27) was added to each well. After 2 days, media were removed and saved at 4°C with added protease inhibitors (P8340, Sigma). Fresh media supplemented with growth factors but without B27 were then added to each well. After an additional 2 days, media were removed and pooled with previously collected media, and additional protease inhibitor was added. The cells were rinsed twice with PBS, and then lysed directly in the well by the addition of 70 μl 2× sample buffer (0.125 M Tris pH 6.8, 20% glycerol, 4% SDS, 0.16% bromophenol blue, 10% 2-mercaptoethanol) and scraping, followed by heating at 95°C for 5 min. All cell lysates were then brought to a total volume of 140 μl with 1× PBS. Supernatants were centrifuged at 14,000×g for 15 min to clear cellular debris, and then were concentrated to ∼30 μl with Amicon 30 kDa filters (UFC503024, Millipore), then mixed with an equal volume of 2× sample buffer and heated at 95°C for 5 min.

Duplicate 4–15% gradient mini-Protean TGX gels (456-1084, Biorad, Hercules, CA) were used to run samples. 40 μl of cell lysate of each genotype or one third of the total volume of concentrated supernatant of each genotype was used. Both gels were then transferred to nitrocellulose membrane and blocked in 3% BSA in TBS plus 0.1% Tween-20 (TBST) for 1 hr at room temperature. Membranes were then incubated overnight with either goat anti-GFRa1 (0.2 μg/ml, AF560, R&D) or goat anti-GFRa2 (0.2 μg/ml, AF613, R&D) in blocking solution overnight at 4°C. Following washes with TBST, membranes were incubated with donkey anti-goat-AP (SC-2022, 1:5000, Santa Cruz Biotechnology, Santa Cruz, CA) in blocking solution for 1 hr at room temperature. After washes, AP was detected with CDP-Star (T2218, Applied Biosystems) and membranes were imaged with a Chemi-Doc system (BioRad).

Following imaging, membranes were stripped with 2× 10 min stripping buffer (0.2 M glycine, 0.1% SDS, 1% Tween-20, pH 2.2), followed by 2× 10 min wash with PBS and 2× 5 min wash with TBST. Membrane was then probed with rabbit anti-β-actin (sc-130656, 1:400, Santa Cruz Biotechnology) and goat anti-rabbit-AP (T2191, 1:5000, Applied Biosystems) following the above procedure, except that all blocking and antibody incubations were performed in 5% milk in TBST.

Western blot densitometry was performed with ImageJ. Three cultures per genotype were analyzed. Densitometry measurements for each antibody were performed on three blots running independent culture samples. Relative quantifications were performed using β-actin in the cell lysates as a measure of total protein per lane, and optical density values for GFRa1 were scaled accordingly. Because an equal proportion of total lysates was run in each lane, total β-actin per cell lysate lane was used as a proxy for cell number, and was therefore used to normalize protein levels in the supernatant lanes (equal proportion of total supernatant volume were run in each lane). Cell lysate and supernatant samples were scaled to wild-type quantifications of respective sample type and reported in arbitrary units. Student's t-test was used to measure significance of differences between genotypes.

Fluorescent images of SC/DRG sections were acquired on a Leica SP5II confocal microscope. DRG explant cultures were imaged on Leica DM5000B microscope. Bright field images were taken using Leica DM5000B microscope.

For histological analysis, at least six sections per specified spinal/DRG level per animal were analyzed. For quantification of genetically labeled neuron number in E18.5 embryos, whole-mount L4/L5 DRGs were imaged and total Tdt⁺ cell number was counted in each DRG. Except for Gfra1;Gfra2 double null animals (n = 2), at least three animals per genotype were examined. N-values for explants are listed in Figure 7—source data 1 and 2. Pixel counts for central projections were generated by counting the number of pixels at each intensity level (0–256) in an outlined immunoreactive area in ImageJ. Background staining was subtracted by counting pixel number of each intensity level in a non-immunoreactive region of the tissue. The minimal intensity level at which two consecutive levels displayed a pixel count of zero was taken as the threshold cut of background fluorescence. Pixel counts of real staining were then calculated by summing the pixel counts for all intensity levels above the defined background level. Column graphs were generated in GraphPad Prism 5. All error bars are ± standard error of the mean (SEM), unless otherwise specified. All statistical analyses were performed using SAS version 9.3 (SAS Inc., Cary, NC). Due to differences in labeling efficiency across litters in 4-HT treated mice, quantification for SC section staining and whole mount DRGs were performed by normalizing to controls within the same litter. For all explant quantifications, GFP⁺ neuron number per 10,000 μm² was calculated for each explant. For Ret^CFP explants, a circle with a radius 200 μm larger than the explant was drawn around the explant in ImageJ, and the number of CFP⁺ axons which crossed the circle was counted.

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

We thank Drs G Bashaw, DD Ginty, M Ma, B Pierchala, and other Luo lab members for helpful comments on the manuscript. We thank Ming Lu for statistical consultation. WL is supported by National Institutes of Health (NIH, R00NS069799 and 1R01NS083702), the Basil O'Connor Starter Scholar Research Award (Grant No. 5-FY12-109 from the March of Dimes Foundation), and the Klingenstein-Simons Fellowship Awards in the Neurosciences. MF is supported by NIH (F31NS086168, T32HD007516, and T32GM07517) and the Hearst Foundation. Work in the Klein lab is supported by the Max-Planck Society.

Animal experimentation: Mice except GDNFlacZ line were raised in a barrier facility in Hill Pavilion, the 618 University of Pennsylvania. All procedures were conducted according to animal protocols 619 approved by Institutional Animal Care and Use Committee (IACUC) of the University of 620 Pennsylvania and National Institutes of Health guidelines. GDNFlacZ mice were raised in 621 accordance with the European Community Council Directive of November 24, 1986622 (86/609/EEC), and approved by the ethics.

© 2015, Fleming et al.

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