extfieldsofstudy: Medicine Biology

text:
Synaptic localization of C9orf72 regulates post-synaptic glutamate receptor 1 levels

A hexanucleotide repeat expansion in a noncoding region of C9orf72 is the most common genetic cause of amyotrophic lateral sclerosis (ALS) and frontotemporal dementia (FTD). Reduction of select or total C9orf72 transcript and protein levels is observed in postmortem C9-ALS/FTD tissue, and loss of C9orf72 orthologues in zebrafish and C. elegans results in motor deficits. However, how the reduction in C9orf72 in ALS and FTD might contribute to the disease process remains poorly understood. It has been shown that C9orf72 interacts and forms a complex with SMCR8 and WDR41, acting as a guanine exchange factor for Rab GTPases. Given the known synaptosomal compartmentalization of C9orf72-interacting Rab GTPases, we hypothesized that C9orf72 localization to synaptosomes would be required for the regulation of Rab GTPases and receptor trafficking. This study combined synaptosomal and post-synaptic density preparations together with a knockout-confirmed monoclonal antibody for C9orf72 to assess the localization and role of C9orf72 in the synaptosomes of mouse forebrains. Here, we found C9orf72 to be localized to both the pre- and post-synaptic compartment, as confirmed by both post-synaptic immunoprecipitation and immunofluorescence labelling. In C9orf72 knockout (C9-KO) mice, we demonstrated that pre-synaptic Rab3a, Rab5, and Rab11 protein levels remained stable compared with wild-type littermates (C9-WT). Strikingly, post-synaptic preparations from C9-KO mouse forebrains demonstrated a complete loss of Smcr8 protein levels, together with a significant downregulation of Rab39b and a concomitant upregulation of GluR1 compared with C9-WT mice. We confirmed the localization of Rab39b downregulation and GluR1 upregulation to the dorsal hippocampus of C9-KO mice by immunofluorescence. These results indicate that C9orf72 is essential for the regulation of post-synaptic receptor levels, and implicates loss of C9orf72 in contributing to synaptic dysfunction and related excitotoxicity in ALS and FTD.

Introduction
Amyotrophic lateral sclerosis (ALS) is a fatal adultonset neurodegenerative disease primarily affecting motor neurons of the motor cortex, brain stem, and spinal cord. With no effective treatment, disease course is typically rapid, resulting in complete paralysis and death within 2-5 years after diagnosis. Hexanucleotide (G4C2) repeat expansions within the first intron of C9orf72 are the most common known genetic cause of both ALS and frontotemporal dementia (FTD) [14,40]. Initial reports on C9orf72 expansions indicated that a length of > 30 was pathogenic; however, there have been several cases where 30-70 repeats do not result in disease, indicating there is no discernible pathological cut-off [17,35,59,60]. As a result, how the expanded G4C2 repeats in C9orf72 cause neurodegeneration in ALS and FTD remains largely uncertain. Three potential pathomechanisms have been proposed to result from the repeat expansions [21,30,52]: (1) RNAmediated toxicity through sequestration of RNA-binding proteins in nuclear repeat RNA foci; (2) accumulation of five dipeptide repeat (DPR) proteins, glycine-alanine (GA), glycine-arginine (GR), proline-alanine (PA), prolinearginine (PR), and glycine-proline (GP), by repeat-associated non-ATG (RAN) translation; and (3) loss of function through C9orf72 haploinsufficiency.
Evidence from human tissues, and cell and animal models has demonstrated that RNA foci are generated in neural cells and the G4C2 repeat structures sequester RNA-binding proteins [1,14,15,17,37,50,63]. In addition, it has been shown that GA, GR, PA, PR, and GP differentially accumulate across different brain regions in ALS/FTD patients [2,3,18,38,39,42,54]. However, evidence has indicated that the distribution of RNA foci and DPRs only show a minor relationship with the severity of neurodegeneration across brain regions, and DPR inclusions in disease are rarely observed in motor neurons at autopsy [12,13,32,33]. Indeed, a recent discovery demonstrated that somatic expansion of the G4C2 repeats does not occur in ALS spinal cord tissues [41]. Interestingly, one group reported an ALS patient presenting with behavioural variant FTD who carried a loss-of-function splice site mutation (c.601 -2A > G) that created a premature stop codon (p.I201fsX235), resulting in reduced C9orf72 mRNA levels in leukocytes relative to control cases [31]. We recently reported a 90-year-old individual carrying 70 G4C2 repeats who was neurologically asymptomatic at autopsy and who had widespread accumulation of RNA foci and DPRs in the brain, but had increased C9orf72 protein levels and no TDP-43 pathology [35,59]. These findings emphasize the importance of assessing the contribution of C9orf72 protein levels to disease mechanism. To date, reduced expression of select or total C9orf72 transcripts [1,6,14,20] or its protein level [57,61] in C9orf72 G4C2 repeat carrier-derived cells or postmortem tissues from C9-ALS/FTD patients have been widely reported. In animal models, knockdown or deletion of C9orf72 orthologues cause motor phenotypes in zebrafish [9] and C. elegans [53], respectively. However, loss of C9orf72 in mice does not induce motor neuron deficits, nor does it produce TDP-43 proteinopathy [24,27]. Collectively, a full understanding of C9orf72 function is needed to elucidate its contribution to the disease mechanism.
Sequence and structure analyses have shown that C9orf72 shares homology with DENN (differentially expressed in normal and neoplastic cells) domain proteins [23,29,66], which are Rab GTPase guanine exchange factors (GEFs) [34,65]. C9orf72 forms a complex with Smcr8 and Wdr41 and can act as a GEF for Rab8 and Rab39b [11,44,62,64]. Given that many Rabs show a synaptosomal distribution [26] and that two studies have shown C9orf72 enrichment at synapses [4,16], we hypothesized that the loss of C9orf72 would lead to alterations in Rab family interactors and glutamatergic receptor levels in synaptosomes. To further understand the biochemical distribution of C9orf72 in the brain, we combined synaptosomal [51] and post-synaptic density preparations [8,10] with a knockout-confirmed monoclonal antibody for C9orf72, which shows both biochemical and immunohistochemical specificity for C9orf72 [28], to assess the distribution of C9orf72 at synaptosomes. Here, we have demonstrated that the mouse ortholog for C9orf72 (31100432021Rik) is localized to both the pre-and post-synapses and is essential for stable Rab39b and GluR1 levels in post-synaptic densities.

Mouse breeding
All animal protocols were conducted in accordance with the Canadian Council on Animal Care and approved by the University of Toronto Animal Care Committee. Knockout C9orf72 mice were obtained through a generous gift from Dr. Don Cleveland (UCSD) and Dr. Clothilde Lagier-Tourenne (UMass). Breeding and genotyping were conducted as previously described [24]. Briefly, C9orf72 heterozygous mice (C57BL/6 background) were crossed, resulting in the production of homozygous C9orf72 KO (C9-KO) and wild-type (C9-WT) littermates. For all biochemistry and immunohistochemical experiments, 3month old mice were empirically selected.

Post-synaptic density fractionation
The fractionation of intact synaptosomes, followed by postsynaptic densities (PSDs), was performed as previously described [8,51] with some adaptations (Fig. 1a). For the characterization of the distribution of protein markers in the PSD fraction of C57BL/6 WT mice, we used three biological replicates (n = 12 mice per replicate). Mice were euthanized with controlled flow CO 2 followed by cervical dislocation. The brains were rapidly removed from the skull and then the cerebellum and brainstem from each brain were discarded, leaving dissected forebrains for each mouse. Mouse forebrains were homogenized with a Dounce homogenizer in ice-cold homogenization buffer (HB; 320 mM sucrose, 5 mM HEPES [pH 7.4]) and then centrifuged at 1000×g for 10 min at 4°C. The resulting pellet was homogenized with RIPA buffer (150 mM NaCl, 50 mM Tris-HCl [pH 7.4], 1% w/v TX-100, 0.5% w/v sodium deoxycholate, 1 mM EDTA, 0.1% SDS) and saved as the post-nuclear/debris fraction (P1). The supernatant (S1) was centrifuged at 13,800×g for 15 min at 4°C. The supernatant (S2) was saved as the cytoplasmic fraction and the pellet (P2) was resuspended in ice-cold HB and then loaded onto a discontinuous Ficoll gradient (13, 9, 5% w/v) prepared in HB and centrifuged at 82,500×g for 120 min at 4°C. Intact synaptosomes (SYN) were retrieved at the boundary between 9 and 13% in the Ficoll gradient and rinsed two times with HB. The resulting pellet was resuspended in cold resuspending buffer (RB; 0.32 M sucrose, 1 mM NaHCO 3 ) and then lysed in an equal volume of lysis buffer 1 (LB1; 1% Triton X-100, 12 mM Tris-HCl [pH 8.1]). The lysed synaptosomes were end-over-end rotated for 15 min at 4°C and centrifuged at 32,800×g for 20 min. The supernatant was saved as the pre-synaptic compartment (S3), whereas the pellet (P3) was resuspended in RB, loaded onto a sucrose gradient (2.0 M, 1.5 M, 1.0 M), and then centrifuged at 201,800×g for 120 min. The crude PSDs were retrieved from the 1.5-2.0 M sucrose interface and resuspended in RB, followed by treatment with equal parts lysis buffer 2 (LB2; 150 mM KCl, 1% TX-100). After end-over-end rotation for 15 min at 4°C, the crude PSD solution was centrifuged at 201,800×g for 4°C. The supernatant was saved as the Triton-KCl treated fraction (S4*), whereas the pellet was resuspended in RIPA buffer and sonicated at < 5 watts to isolate PSDs (P4*).
To allow for the direct comparison of C9-WT versus C9-KO mice (n = 2 per sample; n = 4 biological replicates), the sucrose gradient step was omitted after resuspension of the P3 fraction, as shown in Fig. 1a. We implemented this approach since it was empirically determined that all markers identified in PSDs were present in semi-pure PSDs. For the C9-WT versus C9-KO experiments, P3 was treated directly with RB along with addition of LB2, as described above, followed by end-over-end rotation for 15 min at 4°C. Centrifugation was then performed at 201,800×g for 4°C, where the supernatant was saved as the LB2-treated fraction (S4) and the pellet was resuspended in RIPA buffer and then sonicated (< 5 W) to obtain semi-pure PSDs (P4). Protein concentrations for each fraction were estimated using the bicinchoninic acid assay for electrophoretic loading.

Gel electrophoresis and immunoblotting
Samples were solubilized in sample buffer (3X) and boiled at 95°C for 5 min for immunoblotting. All samples were electrophoresed on 10% (w/v) sodium dodecyl sulphatepolyacrylamide (SDS-PAGE) gels. Separated proteins were transferred to polyvinylidene fluoride (PVDF) membranes. For all immunoblot experiments, membranes were blocked for 1 h at ambient temperature in blocking buffer composed of 5% (w/v) skim milk powder in TBS. PVDF membranes were incubated overnight at 4°C with primary antibodies. A complete list of primary antibodies and the appropriate methodology used is shown in Additional file 1: Table S1. Membranes were then washed with TBS containing 0.05% Tween 20 and incubated for 1 h at ambient temperature with the following secondary antibodies diluted in blocking buffer: either α-mouse horseradish peroxidase (HRP)-conjugated (NA931, GE Healthcare; 1: 5000), α-rabbit HRP-conjugated (NA934, GE Healthcare, 1:5000), or α-goat HRP-conjugated (sc-2433, Santa Cruz, 1:5000) antibody. Visualization of immunoblot labelling was achieved using chemiluminescence with the Western Lighting Plus ECL kit (Perkin Elmer).

Immunoblot image analysis and quantification
Densitometric analysis of immunoblots was performed using the ImageJ distribution Fiji [43]. Protein levels were estimated for each lane by calculating the relative density using an appropriate marker for that fraction (Synaptohysin [Syp] for SYN and S3; PSD-95 for P4). Bar plots were used for visualization of relative protein levels and the statistical analysis comparing C9-WT and C9-KO mice was performed with Prism (v8.0.2, Graph-Pad). For this, multiple t-tests were performed, and the false discovery rate was controlled using the Benjamini, Krieger, and Yekutieli procedure (Q = 1%).

Micrograph image analysis and quantification
For all images, deconvolution of low and high magnification micrographs was performed with 25 iterations and a confidence level of 95% per channel using the Volocity Analysis Suite (v6.3, Perkin Elmer). For GluR1 and Rab39b, fluorescence intensity in the hippocampus of C9-WT versus C9-KO mice was calculated from exported images (n = 3 slices per biological replicate) using Fiji [43]. Qualitative assessment of GluR1 levels indicated increased immunofluorescence across all hippocampal subfields, so the entire hippocampal area was outlined in the fluorescence intensity measurement. For Rab39b, the intensity was measured by outlining the mossy fiber area given the empirically determined distribution in that region alone. Mean fluorescence intensity in each case was calculated by subtracting background intensity from raw intensity and then dividing by the selected hippocampal area. Statistical analyses were performed with Prism (v8.0.2, Graph-Pad), where the mean fluorescent intensity of GluR1 and Rab39b in C9-WT versus C9-KO were contrasted using a paired t-test (p < 0.05).

C9orf72 localizes to both the pre-and post-synapses in murine forebrains
Given that different Rab proteins show unique localization to the pre-and post-synaptic compartment [22,26], we reasoned that C9orf72 and its interacting partner Smcr8 [11,62,64] would be present in post-synaptic densities. To assess this, we performed biochemical fractionation of the pre-and post-synapses in mouse forebrains as described previously [8,51] with some modifications (Fig. 1a). With this approach, we showed enrichment for the pre-synaptic vesicular marker synaptophysin (Syp) in synaptosomes (SYN) and the pre-synaptic compartment (S3), whereas Syp was absent in the post-synaptic fraction (P4*) (Fig. 1b). The established post-synaptic density marker PSD-95 was enriched in the expected fractions (SYN, P3, and P4*) and depleted in the pre-synapses (S3) (Fig. 1b). In addition, postsynaptic receptors GluR1, GluR2, and NMDAR1 were also enriched in P4* (Fig. 1b). We also detected preand post-synaptic TDP-43 in S3 and P4* (Fig. 1b) and found that Rab3a, Rab5, and Rab11 were enriched in the S3 fraction, but depleted in P4* (Fig. 1b). However, Rab39b was found in both the pre-and post-synaptic compartments (Fig. 1b), confirming the differential synaptic distribution of Rab family proteins in the brain [22,26]. Using a knockout-confirmed mouse monoclonal antibody against C9orf72 [28], we detected an~52 kDa band for C9orf72 corresponding to the long isoform of the protein [61] in S3 and P4* (Fig. 1b). We additionally found the C9orf72 interacting partner Smcr8 in S3 and P4* (Fig. 1b), indicating both a pre-and post-synaptic localization of C9orf72 and Smcr8.
To validate C9orf72 presence in the post-synaptic density, we performed immunoprecipitation (IP) of the PSD-95 complex on 12 pooled wild-type mouse forebrains, as previously described [56] with some alterations (Fig. 1a). Both the input (P4*) and IP fractions enriched for the bait protein PSD-95 (Fig. 2a). Post-synaptic receptors GluR1, GluR2, and NMDAR1, in addition to TDP-43, immunoprecipitated with PSD-95 (Fig. 2a), whereas the established contaminants neurofilament light chain (Nefl) and glial fibrillary acidic protein (Gfap) [56] were absent from the PSD IP (Fig. 2a). Notably, C9orf72 was detected in the PSD IP, along with its interacting partners Smcr8 and Rab39b (Fig. 2a). To confirm the brain region localization of synaptic C9orf72, we performed double immunofluorescence labelling with antibody to C9orf72 with either a pre-(Synpr) or postsynaptic (PSD-95) marker on sagittal sections in C9-WT and C9-KO mice. Low magnification images indicated that C9orf72 was present in the forebrain regions that we and others previously identified [16,28], including mossy fiber synapses of the hippocampus (Fig. 2b), the glomerular layer of the olfactory bulb, synapses of the basal ganglia, substantia nigra, and inferior olive, and in the granular layer of the cerebellum (data not shown). We detected partial overlap of C9orf72 with both presynaptic (Synpr) and post-synaptic (PSD-95) markers in the mossy fiber area of the dorsal hippocampus, whereas this synaptic C9orf72 signal was ablated in C9-KO mice (Fig. 2b). Deconvoluted, high magnification images from C9-WT mossy fiber synapses demonstrated partial overlap of C9orf72 with pre-synaptic (Synpr), post-synaptic (PSD-95), and dendritic (MAP 2) proteins (Fig. 2c). Furthermore, high-powered micrographs also confirmed that C9orf72 was partially co-localized with Rab39b in the mossy fiber synapse region of C9-WT mice (Fig. 2c), indicating that C9orf72 shares synaptic localization with previously identified interactors.

Loss of C9orf72 results in decreased Rab39b and increased GluR1 in the hippocampus of C9-KO mice
We asked whether loss of C9orf72 would lead to alterations in post-synaptic Rab39b and receptor protein  levels. For this we performed an immunoblot analysis on semi-pure post-synaptic density preparations (P4; Fig. 1a) from C9-WT and C9-KO mice. We detected stable PSD-95 between C9-WT and C9-KO mice (Fig. 5a), which we verified by densitometry, and then used PSD-95 protein levels to estimate relative abundance for the other markers screened in P4 (Fig. 5b). We observed stable TDP-43, GluR2, and NMDAR1 levels in P4 and undetectable Smcr8 levels in C9-KO preparations (Fig. 5a, b), whereas the relative amount of Rab39b was decreased and GluR1 was increased in P4 from C9-KO versus C9-WT mice (Q < 0.01; Fig. 5a, b). We next assessed the immunofluorescence intensity of Rab39b and GluR1 in 40 μm freefloating sections from C9-WT and C9-KO mouse brains.
There was a concomitant decrease in Rab39b (Fig. 5c) with upregulated GluR1 protein levels (Fig. 5d) in the hippocampus of C9-KO versus C9-WT mice. After quantification, we found a significant decrease in Rab39b fluorescence intensity in the mossy fiber synapse region of C9-KO over C9-WT mice (p < 0.001; Fig. 5e), in agreement with the changes found by immunoblot in P4 (Fig. 5a). We did not detect a significant decrease of Rab39b in the other brain regions where we detected synaptic C9orf72, such as the glomerular layer of the olfactory bulb, synapses of the basal ganglia, substantia nigra, and inferior olive, and in the granular layer of the cerebellum (data not shown). For GluR1, we found a significant increase in fluorescence intensity in the entire dorsal hippocampus (p < 0.05; Fig. 5f), indicating that C9orf72 is required for the proper regulation of hippocampal GluR1 levels in vivo.

Discussion
The elucidation of C9orf72 function remains a crucial research endeavor. The present study explored the effect of in vivo knockout of C9orf72 on pre-and postsynaptic protein expression of C9orf72 interactors such as Smcr8 and Rab family proteins, and on postsynaptic receptor levels. Here we show both biochemical and immunohistochemical evidence that C9orf72 localizes to both the pre-and post-synaptic compartment in the mouse forebrain. Our biochemical results also showed that Smcr8 is present in both the pre-and post-synaptic compartments, and recapitulated previous findings showing that loss of C9orf72 expression results in complete ablation of Smcr8 protein expression in the forebrain (Ugolino, 2016). Our immunoblot studies further showed that Rab3a, Rab5, and Rab11 were localized to the pre-synaptic compartment, and that Rab39b is present in both the pre-and postsynapse. In C9-KO mice, PSDs in the hippocampus demonstrated a concomitant decrease in Rab39b levels with increased GluR1 when compared with C9-WT mice. However, we could not detect any protein level changes between C9-WT and C9-KO mice for GluR2 and NMDAR1 in PSDs. Overall, these results suggest that post-synaptic localization of C9orf72 together with its Rab GTPase Rab39b are important for the regulation of glutamatergic receptor levels in vivo.
Interestingly, loss-of-function RAB39B mutations are causative of both X-linked mental retardation, showing comorbidity with autism spectrum disorder and epilepsy [19,58], and early onset Parkinson's disease with synucleinopathy [58]. RAB39B is important for the coordination of AMPA receptor subunit (GluR1-4) composition and trafficking to the post-synaptic membrane [19,36]. Here, we demonstrated the co-detection of Rab39b with C9orf72 in the hippocampal mossy fiber region of the mouse forebrain, which is similar to a previous study showing RAB39B overlap with C9orf72 in human iPSCderived motor neurons [16]. It has also been shown that knockdown of Rab39b in primary neuron cultures results in increased GluR1 trafficking to dendrites [36]. The current study found that a decrease in Rab39b coincides with increased GluR1 in hippocampal PSDs of C9-KO mice. Hence, this work provides in vivo evidence for a link between postsynaptic C9orf72 and its effect on the protein levels of Rab39b and GluR1 in PSDs.
Although C9-KO mice do not demonstrate an overt behavioural phenotype, we showed that loss of the C9orf72 gene leads to alterations in AMPA subunit levels in the murine hippocampus. AMPA receptor/glutamatergic-associated excitotoxicity is a widely studied disease mechanism underlying ALS [5], FTD [7], and ALS/FTD [49]. In addition to increased GluR1 levels demonstrated in C9-KO mice here, upregulation of GluR1 has been observed in other models, such as cultured neurons from SOD1 G93A [46] and TDP43 A315T [25] transgenic mice that also showed heightened sensitivity to glutamate. Furthermore, increased sensitivity to glutamate has been observed in iPSC-derived neurons from C9-ALS/FTD patients [15,45,47]. Our current findings are in line with a study demonstrating an increase of GluR1 transcript and protein levels with a decreased GluR2 level in iPSC-derived motor neurons from C9-ALS patients [45]. While we did not observe an alteration in GluR2 levels in PSD preparations from C9-KO mice, our results indicate that loss of the C9orf72 gene has specific brain region and synaptic subtype effects on AMPA receptor subunit composition in vivo. Another study showed increased GluR1 and NMDAR1 in human C9-ALS post-mortem motor cortices by immunofluorescence and in PSD preparations [47]. In partial agreement with our study, this study also showed that GluR1 was increased in spinal cord motor neurons from C9-KO mice [47]. More recently NMDAR1 and GluR6/7 levels were shown to be increased in the hippocampus of C9orf72 heterozygous mice [48], indicating that decreasing C9orf72 expression is sufficient to alter glutamatergic receptor levels in vivo. Collectively, the results of these studies and our own converge on glutamatergic excitotoxicity as a potentially important aspect of disease mechanism in both the brain and spinal cord of C9-ALS/FTD.

Conclusion
The present study revealed C9orf72 expression to be an important factor in post-synaptic receptor expression in vivo. We demonstrate a novel localization of C9orf72 to PSDs together with its interactor Smcr8. We also show that loss of C9orf72 in mice results in decreased Rab39b expression and increased GluR1 levels in the hippocampus. Future studies will assess the functional association between post-synaptic C9orf72 and Rab39b protein levels on AMPA receptor trafficking. A better understanding of glutamatergic excitotoxicity in C9-ALS/FTD and in non-C9 ALS/FTD may lead to novel therapeutic development.

Additional file
Additional file 1: Table S1. Primary antibodies used in the current study. Figure S1. Rab39b and GluR1 protein levels are unchanged in S1 fraction between C9-WT and C9-KO mice. Bar plots of mean S1 immunoblot band densities for (GAPDH); Ras-related protein Rab39b; Glutamate receptor 1 (GluR1). y-axis = relative density; error bars = standard deviation. All pairwise comparisons are not significant.


extfieldsofstudy: Biology Medicine

s2fieldsofstudy: Biology Medicine

text:
Roles of Primary Cilia in the Developing Brain

Essential to development, primary cilia are microtubule-based cellular organelles that protrude from the surface of cells. Acting as cellular antenna, primary cilia play central roles in transducing or regulating several signaling pathways, including Sonic hedgehog (Shh) and Wnt signaling. Defects in primary cilia contribute to a group of syndromic disorders known as “ciliopathies” and can adversely affect development of the brain and other essential organs, including the kidneys, eyes, and liver. The molecular mechanisms of how defective primary cilia contribute to neurological defects, however, remain poorly understood. In this mini review, we summarize recent advances in understanding of the interactions between primary cilia and signaling pathways essential to cellular homeostasis and brain development.

INTRODUCTION
Primary cilia are microtubule-based cellular organelles that aid in sensing and transducing environmental signals during development (Goetz and Anderson, 2010): primary cilia transduce or regulate several signaling pathways, including Sonic hedgehog (Shh) and Wnt signaling (Corbit et al., 2005;Rohatgi et al., 2007;Gerdes and Katsanis, 2008;Goetz et al., 2009). This antenna-like structure was first observed using electron microscopy at the lumen of kidney tubules in 1898 and has since been intensively studied (Zimmermann, 1898). Emanating from the apical surface of basal body, the axoneme structure of primary cilium comprises a radial array of nine microtubule doublets lacking a central pair (9+0 structure) ( Figure 1A; Davenport and Yoder, 2005). At the base of primary cilium, protein entry and exit are regulated via a transition fiber that anchors the axoneme to the ciliary membrane, compartmentalizing this distinct organelle from the cytosol while remaining continuous with the plasma membrane (Gherman et al., 2006;Marshall, 2008;Reiter et al., 2012).
Leading to defective primary cilia, mutations in genes necessary for ciliogenesis and ciliary structure and function are known to cause a group of human genetic disorders described as "ciliopathies." To date, 187 mutated genes in 35 known ciliopathies and 241 ciliopathyassociated genes essential to ciliary structure and function that could potentially be causative for ciliopathies have been documented (Reiter and Leroux, 2017). These ciliopathies affect the body's essential organs, including the kidneys, eyes, brain, liver, and skeleton, during development and tend to share clinical phenotypes (Hildebrandt et al., 2011): for example, as primary cilia are critical to development of the central nervous system (CNS), many ciliopathies, such as Joubert syndrome (JBTS), Meckel syndrome (MKS), and orofaciodigital syndrome (OFD), commonly exhibit neurological defects of CNS malformation, intellectual disability, ataxia, and retina dystrophy (Valente et al., 2014). Here, we summarize our recent understanding of the interactions between primary cilia and several signaling pathways essential for cellular homeostasis and for brain development.  (Rohatgi et al., 2007;Tukachinsky et al., 2010;Sasai and Briscoe, 2012). This initiates the activation of Gli transcriptional factors ( Figure 1B; Alcedo et al., 1996): Gli regulates the transcription of target genes that regulate the Shh signaling (e.g., Ptch1, Gli1), proliferation (e.g., cyclin D1, MYC), and apoptosis (e.g., Bcl-2) (Scales and de Sauvage, 2009). Shh, Ptch, Smo, and Gli are expressed in the developing and adult brain (Echelard et al., 1993;Ekker et al., 1995).

SIGNALING PATHWAYS AND
Sonic hedgehog signaling is critical to spatial patterning of the neuroepithelium, cellular identity in CNS, axonal guidance, wiring of the neural network, and neuronal activity. As Shh signaling-related proteins are located within primary cilium, Shh signaling is not properly achieved in mice with defective cilia, resulting in several defects in brain development such as defective neural patterning, cerebellar hypoplasia, and defective hippocampal neurogenesis (reviewed in Ferent and Traiffort, 2015). For example, mice with mutation in Kinesin family member 3A (Kif3a), which is important for ciliogenesis, show reduced Gli1 expression, thereby leading to defects in hippocampal neurogenesis and formation of neural stem cells . Stumpy mutant mice exhibit defective ciliogenesis leading to prenatal hydrocephalus and severe polycystic kidney disease. The Stumpy gene, located on mouse chromosome 7, encodes B9 protein domain 2 (B9D2), which is localized with basal bodies along ciliary axonemes and appears to play a role in ciliogenesis in association with intraflagellar transport proteins (IFTs) (Town et al., 2008). Conditional Stumpy mutant mice (driven by Nestin-Cre) have been found to show abrogated Shh signaling and Gli processing in the hippocampus .

Wnt Signaling
Wnt signaling is another developmental signaling pathway mediated by primary cilia (Figure 1C; May-Simera and Kelley, 2012). Wnt signaling can be divided into (1) β-catenin dependent (canonical) and (2) β-catenin independent (non-canonical) signaling (reviewed in Berbari et al., 2009;Oh and Katsanis, 2013). Several studies have shown that perturbation of ciliary genes aberrantly activates canonical Wnt signaling and disrupts non-canonical Wnt signaling (Lin et al., 2003;Cano et al., 2004;Ross et al., 2005;Simons et al., 2005;Gerdes et al., 2007;Corbit et al., 2008). In particular, Inversin, which is localized in cilium, mediates the proteasomal degradation of Dishevelled (Dvl) to regulate both canonical and non-canonical Wnt signaling (Simons et al., 2005). Perturbations in several ciliopathy-related genes have been found to elicit de-regulated canonical Wnt signaling in fibroblasts and in the forebrain (Willaredt et al., 2008;Abdelhamed et al., 2013;Wheway et al., 2013). Also, ciliary ablation in adult-born dentate granule cells in the hippocampus has been shown to aberrantly activate canonical Wnt signaling, concomitant with severe defects in dendritic refinement and synapse formation (Kumamoto et al., 2012). Additionally, researchers have recently demonstrated that defective neuronal ciliogenesis caused by hyperactivating mutation in MTOR elicits abnormal activation of canonical Wnt signaling and inactivation of non-canonical Wnt signaling, resulting in defective neuronal migration due to disrupted neuronal polarization (Park et al., 2018). Notwithstanding, several studies have reported that ciliary dysfunction does not affect Wnt signaling, as evidenced by constant activity of Wnt signaling reporter and expression of Wnt target genes (Huang and Schier, 2009;Ocbina et al., 2009). Interestingly, however, deregulated canonical Wnt signaling via deletion of adenomatous polyposis coli (APC) was found to cause a loss of primary cilia in association with radial progenitor malformation in the neocortex (Nakagawa et al., 2017). In these regards, the interaction between primary cilia and Wnt signaling may be context-dependent, and more thorough studies will be necessary to elucidate it.

MTOR Signaling
MTOR is a serine/threonine kinase essential for protein translation, lipid synthesis, and autophagy, and plays an essential role in neural differentiation, neuronal migration, and synaptic formation, all of which are crucial to brain development. Disruption of MTOR signaling has been documented in numerous pathological conditions, including cancer, neurological disorder, and metabolic disorder (Lipton and Sahin, 2014;Saxton and Sabatini, 2017) Polycystic kidney disease (PKD) patients with inherited mutations in ciliary genes show deregulation of the MTOR signaling pathway (Shillingford et al., 2006;Wahl et al., 2006). In particular, polycystin-1, which is recurrently mutated in PKD, has been shown to inhibit MTOR through interaction with TSC2, a component of the TSC complex that inhibits MTOR (Shillingford et al., 2006). Studies have also identified that, by bending stimulus via fluid flow, primary cilia downregulate AMPK-MTOR signaling, which, in turn, induces autophagy to control cell size via LKB1 localized at primary cilia in the kidneys (Boehlke et al., 2010;Orhon et al., 2016). Recently, research has demonstrated that, during brain development, mice with defective cilia present abnormal increases in MTOR signaling, leading to enlarged apical domains of radial glial cells (RGCs) and subsequent dilatation of brain ventricles (Foerster et al., 2017). Conversely, in TSC1-or TSC2-null fibroblasts, TSC1 and TSC2 have been found to positively regulate ciliogenesis without using the TSC-MTOR signaling axis (i.e., rapamycin-insensitive) (Hartman et al., 2009). Nevertheless, a recent independent study which also performed with TSC1-or TSC2-null fibroblasts reported different ciliary phenotypes marked by a longer ciliary length in TSC1-null cells and a shorter ciliary length in TSC2-null cells, and there was a discrepancy between the two cell types in that only the elongated ciliary phenotype in TSC1-null cells could be rescued by rapamycin treatment (Rosengren et al., 2018). In focal malformations of cortical developments (FMCDs), such as hemimegalencephaly (HME) and focal cortical dysplasia (FCD), which are highly associated with intractable epilepsy and autism-spectrum disorders, brain somatic activating mutations in MTOR eliciting blockage of autophagy have been described as disrupting neuronal ciliogenesis in brain tissues from FMCD patients (Wegiel et al., 2010;Lim and Crino, 2013;Park et al., 2018; Figures 2A,B). Given the complex reciprocal interaction between primary cilia and MTOR signaling, further mechanism studies are yet needed.

Autophagy
Autophagy is a cellular degradative process by which a cell gains nutrients to maintain homeostasis (Mizushima, 2007). Two independent studies have provided evidence of links between ciliogenesis and autophagy: In serum-nutrient conditions, basal autophagy degrades the ciliary protein IFT20, while OFD1 protein at centriolar satellites inhibits ciliogenesis. Upon serumwithdrawal, the centriolar satellite pool of OFD1 is degraded by inducible autophagy, while IFT20 initiates ciliary trafficking, leading to ciliogenesis (Pampliega et al., 2013;Tang et al., 2013). Cilia-mediated Shh signaling has been found to assemble several proteins needed for autophagy at the periciliary region, thus activating Shh signaling induces autophagy flux. Interestingly, one study described defects in serum-withdrawal-induced autophagy and biogenesis of autophagosome among Ift20-or Ift88-compromised cells (Pampliega et al., 2013). In line with this, others have found that treatment of Shh to culturedhippocampal neurons upregulates several autophagy-related genes and enhances autophagy (Petralia et al., 2013).
Meanwhile, dysregulated ciliogenesis in relation to defective autophagy has been described in several diseases. Previous studies have reported suppressed autophagic flux in ciliary dysfunctional PKD mouse models and impairment of autophagosome formation in cells derived from PKD patients (Belibi et al., 2011;Zhu et al., 2017). Interestingly, Hürthle cells found in thyroid cancer were found to show defective ciliogenesis, and this defect in Hürthle cells eliciting high basal autophagic flux was restored by autophagy inhibition (Lee et al., 2016). Dysregulated ciliogenesis has also been reported in Huntington's disease (HD), for which defective autophagy has been well recognized (Keryer et al., 2011;Kaliszewski et al., 2015). Although Huntingtin, which is mutated in HD, is known to regulate autophagy, it also participates in trafficking pericentriolar material 1 (PCM1) to the centrosome. When Huntingtin is mutated, aberrant accumulation of PCM1 occurs, which, in turn, causes dysregulated ciliogenesis (Keryer et al., 2011;Rui et al., 2015). Further studies, however, are required to elucidate the direct interaction between ciliary abnormality and defective autophagy in HD. Finally, as stated in the previous section on MTOR signaling, defective autophagy and resultant disruption of ciliogenesis have been demonstrated in FMCDs (Park et al., 2018). Brain somatic activating mutations in MTOR, which are causative for FMCDs, block autophagy, resulting in defective neuronal ciliogenesis due to aberrant accumulation of OFD1 ( Figure 2C; Park et al., 2018). Although defective autophagy has been implicated in many neurodevelopmental and neurodegenerative disorders, much more studies are needed to elucidate the pathophysiological role of primary cilia in these disorders (Levine and Kroemer, 2008;Lee et al., 2013;Marsh and Dragich, 2018).

Patterning and Morphogenesis of the Forebrain
One of the well-established developmental functions of primary cilia is to control forebrain patterning, which is severely defected in human ciliopathies (Vogel et al., 2012). Patterning in the neural tube of the CNS occurs through progressive subdivisions along the dorsal to ventral and the rostral to caudal axes (Dessaud et al., 2008). Cobblestone mice (Ift88 cbs/cbs ), which show severe disorganization of the telencephalon, have been found to exhibit an ambiguous dorsal-ventral forebrain boundary and dorsal telencephalic-diencephalic boundary, concomitant with increased levels of full-length Gli3 (Willaredt et al., 2008). Meanwhile, mice lacking primary cilia as a result of mutation in the ciliopathy gene Rpgrip1l exhibited defective phenotypes associated with MKS and JBTS. In these mice, a mislocalized olfactory bulb-like structure and ambiguous dorsalventral patterning were observed, both of which were rescued upon introduction of constitutively active repressor form of Gli3 (Besse et al., 2011). Other mice with defective cilia through loss of Ttc21b (encoding retrograde Ift139) have also been shown to display defects in dorsal-ventral patterning and rostralcaudal patterning. Researchers have reported that Ttc21b-null mice exhibit normal anterograde transport that permits entry of Shh signaling components into cilium but partially defective retrograde transport. Overactivation of GLI2 and GLI3A in these mice could accounts for activation of Shh signaling and abnormal patterning, suggesting a periciliary role for retrograde IFT in GLI3 processing and GLI2 activity (Huangfu and Anderson, 2005;May et al., 2005;Tran et al., 2008;Stottmann et al., 2009). Crossing Ttc21b-null mice with mice heterozygous for a null allele of Shh to reduce levels of Shh ligand was found to partially rescue their defective phenotypes (Tran et al., 2008;Stottmann et al., 2009). This paradoxical effect of retrograde IFT mutants on Shh signaling (i.e., activation of Shh signaling) was well reviewed by Pal and Mukhopadhyay (2015) and Bangs and Anderson (2017). In line with the importance of primary cilia in cortical patterning, mice with mutant Slb (encoding anterograde Ift172) that possess very short axonemes with no visible microtubules FIGURE 2 | Defective ciliogenesis due to brain somatic mutations in MTOR accounts for cortical dyslamination in FMCDs. (A) Representative brain MRIs of patients with FMCDs, including hemimegalencephaly (HME) and focal cortical dysplasia (FCD) type II. Arrows indicate the affected region of the brain. Adapted from Park et al. (2018), with permission. (B) Immunostaining for Arl13b, a marker for primary cilia, and NeuN, a marker for neurons, with DAPI co-staining in brain tissue from FMCD patients. While primary cilium normally forms at each neuron in brain tissue from non-FMCD, FMCD patients with brain somatic mutations in MTOR exhibit defective neuronal ciliogenesis. Scale bars, 30 µm. Adapted from Park et al. (2018), with permission. (C) Autophagic degradation of the centriolar satellite pool of OFD1 induces primary ciliogenesis. However, brain somatic activating mutation in MTOR, which blocks autophagy, disrupts ciliogenesis in brain tissues with FMCDs. (D) Cortical dyslamination with ciliary defective dysmorphic neurons in the cerebral cortex of a FMCD patient. Somatic activating mutations in MTOR, causative for FMCDs, disrupt neuronal ciliogenesis through blockage of autophagy, resulting in cortical dyslamination.
in their cilia have also been found to exhibit a global brainpatterning defect along the dorsal-ventral and rostral-caudal axes (Gorivodsky et al., 2009). Collectively, these studies suggest the importance of Gli3 processing, which could be mediated by primary cilia, in forebrain patterning.
The establishment of polarized radial glial scaffold formation during cortical development has been shown to be regulated by Arl13b, a small GTPase enriched in primary cilia (Higginbotham et al., 2013). Arl13b mutant mice (Arl13b hnn/hnn ), which display defective cilia, surprisingly exhibit dramatic inversion of the apicobasal polarity of the radial glial scaffold, which, in turn, changes the location of the progenitor zone to the outer part of the cortex. This defect is also observed when Arl13b is removed just before the formation of radial glia from neuroepithelial cells using Foxg1-driven Cre. However, Arl13b removal after establishment of the radial glial scaffold using Nestin-or GFAP-driven Cre has not been found to generate this intriguing phenotype, suggesting that the role of Arl13b in polarization is temporally restricted to the period during which the radial glial scaffold develops (Higginbotham et al., 2013). This reversed apicobasal polarity seen in Arl13b mutant mice has not been reported in mice deficient of other ciliary genes. In line with this temporally restricted role of a ciliary gene during development, a recent study performed conditional knockout (KO) of Kif3a, Ift88, and Ttc21b in a series of specific spatiotemporal domains. Intriguingly, the observed neurological defects, including forebrain expansion, cortical malformations, impaired olfactory bulb development, and ventriculomegaly, were significantly different across four different Cre transgenic alleles (Foxg1-Cre, Emx1-Cre, Wnt1-Cre, and AP2-Cre), indicating that the roles of cilia in cortical development are discretely spatiotemporal. For example, the deletion of Ift88 at earlier stages of development, driven by Foxg1-Cre, was found to increase brain size; however, when deleted at later stages, driven by Emx1-Cre, there were no significant differences in brain size (Snedeker et al., 2017). Taken together, these experimental studies of perturbations of primary cilia in mice support the crucial role of primary cilia in forebrain development, which, when disrupted, lead to severe morphological malformations in the brain.

Neuronal Migration
During brain cortical development, newly born neurons follow one of two main migratory routes to reach their final destination, known as radial and tangential migration (Hatten, 1999). The role of primary cilia in radially migrating pyramidal neurons remains unclear. In a previous study, cerebral cortical lamination was not apparently affected in Arl13b-conditional KO mice driven by Nex-Cre or in Stumpy-, Kif3a-, and Ift88-conditional KO mice driven by Nestin-Cre (Arellano et al., 2012;Higginbotham et al., 2012;Tong et al., 2014). However, a recent in vivo systemic study revealed that genes linked to ciliopathies affect cortical development, including neuronal radial migration, neural progenitor development, neuronal differentiation, and early neuronal connectivity (Guo et al., 2015). In this study, knockdown of 17 ciliopathy genes in the developing neocortex resulted in delayed neuronal radial migration in association with transient multipolar stage, multipolar-to-bipolar transition, and glial-guided radial migration (Guo et al., 2015). Meanwhile, others have suggested that the defective phenotypes seen in ciliary conditional KO mice driven by Nestin-Cre, the expression of which is broad and is also seen in non-RGCs at later developmental stages, may be partly attributable to non-cellautonomous effects (Foerster et al., 2017;Chen et al., 2018). Accordingly, the cell-autonomous function of primary cilia in neural stem cells at the late developmental stage was investigated via acute knockdown of ciliary genes (Kif3a or Ift88) in the developing cortex, which resulted in defective neuronal differentiation and migration; this was accompanied by delays in neural stem cell cycle progression and failures in interkinetic nuclear migration (Chen et al., 2018). In FMCDs, the molecular mechanism of how somatic mutations in MTOR lead to focal cortical dyslamination appears to be related to defective neuronal ciliogenesis by MTOR mutation, which affects the multipolar to bipolar transition essential for neuronal radial migration (Figures 2C,D; Park et al., 2018).
Interneurons originating from the medial ganglionic eminence migrate along a tangential path in the cortical plate, either at the marginal zone or intermediate zone. These neurons change their orientation from a tangential to radial path to colonize the cortical plate and to differentiate into GABAergic neurons (Guo and Anton, 2014). Recent studies have demonstrated that disruption of primary cilia at interneurons affects the interneuronal migratory process, suggesting the importance of primary cilia in neuronal migration (Baudoin et al., 2012;Higginbotham et al., 2012). In a previous study, mice with conditional KO of Arl13b genes driven by Dlx5/6-Cre showed disruption of interneuron placement at Arl13b-deleted cortices caused by defects in interneuron migration and branching. However, when the Arl13b gene was disrupted in post-migratory interneurons driven by PV-Cre, there was no significant difference in the morphology of dendritic arbors during neuronal differentiation. Such defective interneuron migration may be a potential pathophysiological mechanism underlying neurological defects in JBTS with Arl13b mutations (Higginbotham et al., 2012). Meanwhile, a recent study showed that disruption of cilia in interneurons using Nkx2.1-Cre to ablate Ift88 or Kif3a genes led to defective migration of interneurons, marked by an aberrant accumulation of interneurons at their first tangential path, not radial streams in the cortical plate, as well as abnormal positioning and density of interneurons in the postnatal cortex (Baudoin et al., 2012). Taken together, these studies emphasize that primary cilia in immature interneurons are crucial to interneuron migration in the developing cortex and the underlying mechanisms thereof (Baudoin et al., 2012). Nevertheless, the role of primary cilia in neuronal migration needs further study, especially in regards to which other ciliamediated signaling pathways are involved. These future studies on neuronal migration should shed light on the pathophysiology of ciliopathies presenting with cognitive deficits, which may be attributable to defective neuronal migration and disrupted neural connectivity.

Cerebellum Development
Ataxia due to cerebellar malformation is one major symptom of ciliopathies, particularly JBTS. During development of the cerebellum, Purkinje cells secrete Shh, which regulates the proliferation of cerebellar neuronal precursor (Wechsler-Reya and Scott, 1999;Haldipur et al., 2012): Shh signaling mediated by primary cilia occurs in late embryonic stages of cerebellum development, during which Shh is required for expansion of the granule neuron precursor population, but not for the subsequent differentiation of these cells (Lewis et al., 2004). Primary cilia control cerebellar morphogenesis by promoting the expansion of the granule progenitor pool. Loss of either Ift88 or Kif3a causes severe cerebellar hypoplasia and inhibits the expansion of the granule cell progenitor population. Although Ift88 and Kif3a are not required for the specification and differentiation of cerebellar cell types, they affect granular cell progenitor proliferation. Researches have shown that the expression of Gli1 at the external granule cell layer in Kif3a mutant mice is lower than that in controls (Chizhikov et al., 2007;Spassky et al., 2008). These studies demonstrate that ablation of primary cilia impairs Shh signaling, which regulates the development of the cerebellum, particularly expansion of cerebellar progenitors. This mechanism could explain the hypoplasia of the cerebellum and cerebellar ataxia seen in patients with ciliopathies such as JBTS.

Learning and Memory
Patients with ciliopathies commonly present with intellectual disability (ID) as a neurological deficit. Although there are not enough studies to explain the mechanism underlying ID as a result of defects in primary cilia, several studies have shown that depletion of primary cilia leads to hippocampal-dependent learning and memory deficits Han et al., 2008;Amador-Arjona et al., 2011). Hippocampal neurogenesis plays a role in fear conditioning, recognition, spatial memory, and pattern separation (Cameron and Glover, 2015). Also, Shh signaling mediated by primary cilia controls the proliferation of neural progenitor cells (NPCs). As evidence thereof, Emx1-Credependent Shh-conditional KO mice were found to show smaller dorsal telencephalons at E18.5 and decreased proliferation and increased apoptosis of neural stem cells (NSCs) and NPCs in the dorsal pallium (Komada et al., 2008). Meanwhile, Nestin-Cre-and hGFAP-Cre-dependent Smo KO mice exhibited smaller postnatal dentate gyruses and fewer proliferating cells than those in controls (Machold et al., 2003;Han et al., 2008). In Kif3a mutant and Ift88 mutant mice, neuronal proliferation in the subgranular zone (SGZ) and subventricular zone (SVZ) was decreased, and Gli1 expression was abolished Tong et al., 2014;Gazea et al., 2016) These demonstrate that Shh signaling is involved in the proliferation and survival of NSCs and NPCs. Hypomorphic Ift88-mutant mice, in which Ift88 mRNA and protein are reduced by 70-80%, showed disorganization of the midbrain dopominergic neuron (mDA) progenitors domain, and the number of mDA neurons was severly reduced (Gazea et al., 2016). Considering that primary cilia regulate neurogenesis by mediating Shh signaling, defects in learning and memory and ID might be caused by decreased neurogenesis due to aberrant Shh signaling. Expanding on the above research, additional studies are needed to determine how defective neuronal cilia lead to learning and memory impairment and ID.

CONCLUDING REMARKS
In this review, we briefly summarized the interaction between primary cilia and several signaling pathways, including Shh, Wnt, MTOR, and autophagy, all of which are essential to cellular homeostasis. Given that neurological disorders are typical symptoms of ciliopathies, the importance of primary cilia during brain development cannot be emphasized enough. At present, however, we are unable to fully understand the underlying mechanisms by which defective neuronal cilia contribute the pathogenesis of neurological defects in ciliopathies. Future studies of neuronal cilia in relation to signaling pathways may provide better understanding of the functions of neuronal cilia and may help to develop novel treatments for ciliopathies by targeting neuronal cilia-mediated signaling pathways. Compared with considerable studies on neuronal cilia, there is a lack of research on the role of primary cilia in glial cells, another major component of the brain. Glial cells, such astrocytes and oligodendrocytes, also possess a single primary cilium Bishop et al., 2007;Kasahara et al., 2014). Recently, studies reported defective ciliogenesis in glioblastoma (GBM), a high-grade astrocytic malignancy, and demonstrated the potential role of primary cilia on GBM progression (Moser et al., 2014;Hoang-Minh et al., 2016a,b;Loskutov et al., 2018). Further studies on the role of primary cilia in glial cells, as well as neurons, will advance our understanding of the role of primary cilia in brain development.
To help achieve a better understanding of human ciliopathies resulting from mutations in ciliary genes, CRISPR/Cas9 genome editing of identified mutations in human ciliopathies or 3D culture of brain organoids derived from pluripotent stem cells from ciliopathy patients can be used to generate more pathophysiologically relevant models for ciliopathy, thereby overcoming the gap between animal models and human subjects. Also, increasing studies on interactions between primary cilia and other signaling pathways may uncover a previously unknown role for cilia in the pathogenesis of human neurological diseases that are not considered classical ciliopathies, such as FMCD, brain cancer (medulloblastoma and GBM), and neurodegenerative diseases (Alzheimer's disease and HD) (Sotthibundhu et al., 2009;Keryer et al., 2011;Eguether and Hahne, 2018). Future studies of these disorders may help us to further the molecular basis of human diseases characterized by defects in primary cilia.

AUTHOR CONTRIBUTIONS
SP, HJ, and JL conceived the idea and wrote the manuscript.

FUNDING
This work was supported by grants from the Suh Kyungbae Foundation (to JL); Korean Health Technology R&D Project, Ministry of Health and Welfare, Republic of Korea (H15C3143 and H16C0415 to JL).


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Genetics and functions of the retinoic acid pathway, with special emphasis on the eye

Retinoic acid (RA) is a potent morphogen required for embryonic development. RA is formed in a multistep process from vitamin A (retinol); RA acts in a paracrine fashion to shape the developing eye and is essential for normal optic vesicle and anterior segment formation. Perturbation in RA-signaling can result in severe ocular developmental diseases—including microphthalmia, anophthalmia, and coloboma. RA-signaling is also essential for embryonic development and life, as indicated by the significant consequences of mutations in genes involved in RA-signaling. The requirement of RA-signaling for normal development is further supported by the manifestation of severe pathologies in animal models of RA deficiency—such as ventral lens rotation, failure of optic cup formation, and embryonic and postnatal lethality. In this review, we summarize RA-signaling, recent advances in our understanding of this pathway in eye development, and the requirement of RA-signaling for embryonic development (e.g., organogenesis and limb bud development) and life.

Background
For human health, the importance of retinol, also known as vitamin A, has been known since ancient times, when the practice of squeezing liver juice into the eye was used as a treatment for night blindness [1]. The link between night blindness (nyctalopia) and nutrition was first described by Hippocrates during the 4th century BC, when he recommended eating raw liver in combination with honey as a cure [2].
The idea that certain foods possessed curative properties was understood for much of human history. However, it was not until much later that a series of controlled experiments (i.e., human dietary supplementation and animal models during the early twentieth century) allowed scientists to investigate how removal of certain factors from the diet could cause debilitating illnesses and death [3][4][5]. Biochemical experiments in vertebrate models subsequently revealed that retinol was the active compound involved in cell growth and development, along with its precursors and metabolites [6].
The aldehyde derivative of retinol, 11-cis-retinal, is required for vision [reviewed in [7]]. All-trans-retinoic acid (ATRA), the acid derivative of retinol, is able to prevent developmental defects in vitamin A-deficient animals [8]. The demonstration that retinoic acid (RA) could not be converted back to retinol in vivo led to the conclusion that RA was a necessary nutrient involved in cell growth and development [9]. Ultimately, a set of compounds and their metabolites with biological functions similar to retinol were termed "retinoids" [10].
Although the chemical structures of retinoids were identified in the early 1900s, little was known about the mechanisms by which these small lipophilic molecules exerted their biological effects [6]. A short time later, studies performed in vitamin A-deficient rats revealed that retinol supplementation stimulated RNA synthesis in intestinal cells [11]. Biochemical studies subsequently identified serum, membrane, and cytosolic proteins that were essential for retinol transport, uptake, and metabolism [12]. Examples include retinol-binding protein 4 (RBP4), stimulated-by-retinoic-acid 6 (STRA6) membrane receptor, and the cellular retinol-binding protein (CRBP) family, also known as RBP1 and RBP2 [12].
Experiments carried out in chick and mouse embryos identified RA as the active metabolite of vitamin A that possessed the ability to regulate cellular differentiation and proliferation, as well as pattern formation during embryogenesis [reviewed in [13]]. At the molecular level, the action of RA is mediated by two distinct classes of proteins: (i) a family of nuclear receptors comprising RA receptors (RARs) and retinoid X receptors (RXRs) that regulate gene transcription in a ligand-dependent fashion and (ii) a family of cytosolic proteins called cellular retinoic acid-binding proteins (CRABP1 and CRABP2) which facilitate cellular RA uptake and nuclear transfer [14]. These studies provided a link between the chemical structure of retinoids and their biological action.
Although the developmental role of RA has been extensively studied in model organisms [reviewed in [13]], little is known about the exact role of RA in human development. Currently, almost all our molecular understanding about the pathogenesis of RA deficiency is based on either vertebrate animal knockout models (in which genes encoding proteins involved in RA synthesis or degradation are selectively inactivated) or experiments involving rodents fed diets deficient in vitamin A. Such studies have shown RA to be involved in early axial and central nervous system patterning, neurogenesis, regulation of limb bud development, and organogenesis [ [3] and reviewed in [13,14]]. To date, none of these studies has been systematically validated in humans due to ethical concerns and the difficulty of performing such clinical experiments. Nevertheless, based on observational studies in humans consuming vitamin A-deficient diets, it is well established that vitamin A is required in humans (even into adulthood) because it regulates fertility, maintains normal vision, inhibits neoplastic growth, and prevents neurodegenerative diseases [15].
With the recent publication of whole-exome sequencing (WES) data from ≈ 140,000 individuals by the Genome Aggregation Database (gnomAD) [16], it is now possible to investigate genetic intolerance to protein-truncated variants (PTVs) in a large population, i.e., to detect genes that are essential for human development. In this review, we first provide an overview of canonical and non-canonical RA metabolism (i.e., synthesis and catabolism) and the mechanism of RA target gene regulation. We then provide an update on the role of RA-signaling in eye development in mouse and zebrafish and discuss the ocular diseases in humans who have mutations in genes involved in the RAsignaling pathway-such as microphthalmia, coloboma, and anophthalmia. Finally, we take advantage of population-level variation databases to identify which genes involved in the RA pathway display loss-of-function intolerance, thus indicating their requirement for human development and life.

Main text
Retinoic acid synthesis, catabolism, and gene regulation This section describes the components of the retinoic acid signaling pathway including cellular uptake of retinol, conversion of retinal to retinaldehyde, retinaldehyde oxidation to RA, RA degradation, and target gene activation (Fig. 1).

Canonical pathway of RA synthesis
Early studies revealed that retinoids could not be synthesized de novo in most animals [reviewed in [13]]. Hence, the major source of retinoids during embryonic and fetal development is through placental transfer of maternal retinol. Postnatally, retinoids are primarily derived from the dietary intake of (i) carotenoids (such as β-carotene) contained in plant pigments and (ii) retinyl esters from animal sources, such as fish-liver oils, eggs, milk, and butter. Following ingestion, retinyl esters are hydrolyzed to retinol by intestinal mucosal enzymes, whereas carotenoids are cleaved into retinal and subsequently reduced to retinol or oxidized to RA. Retinol homeostasis is tightly regulated. As such, much of the synthesized retinol is converted back into retinyl esters for storage in liver hepatocytes and stellate cells. When needed, these esters are cleaved and released into the bloodstream as retinol [17,18]. Upon release into the bloodstream, retinol is bound by retinol-binding protein 4 (RBP4).
Cells can take up the retinol-RBP4 complex via transmembrane receptor protein stimulated by retinoic acid 6 (STRA6), the product of the RA-inducible mouse gene Stra6 (or human STRA6 gene) [19]. The complex tissuespecific expression pattern of this gene during development influences which tissues are able to take up retinol [20]. Once inside the cell, two sequential reactions are required to transform retinol into retinaldehyde and RA (Fig. 1).
The first reaction is mediated by two classes of enzymes: (i) cytosolic alcohol dehydrogenases (ADHs) that belong to the medium-chain dehydrogenase/reductase family and/or (ii) microsomal retinol dehydrogenases (RDHs) that belong to the short-chain dehydrogenase/ reductase family [21]. Initial studies in mice indicated that this reaction was catalyzed by ADH7 in the embryo [22,23]; however, tissue-specific RDH10 is now believed to play the most important role in RA synthesis during development because mice expressing mutant Rdh10 (RDH10 trex ) (that lacks the ability to convert retinol to retinal) display embryonic lethality [24]. Some degree of RA activity persists in mice expressing RDH10 trex (revealed by limited RARE-lacZ reporter activity at E9.5), indicating that other enzymes (such as ADH7) are able to generate retinaldehyde, albeit at lower levels (that do not support embryonic development) [24]. In addition, transgenic suppression of ADH5 (an enzyme ubiquitously expressed in embryo and adult) or of tissuespecific ADH1 and ADH7 revealed that ADH enzymes may have a role in controlling removal of excess retinol, rather than participating in RA synthesis per se [25].
The second reaction is oxidation of retinaldehyde to RA. This is catalyzed by three aldehyde dehydrogenases: ALDH1A1, ALDH1A2, and ALDH1A3 which are encoded (respectively) by Aldh1a1, Aldh1a2, and Aldh1a3 (respectively) in mice, or by ALDH1A1, ALDH1A2, and ALDH1A3 in humans. Each ALDH displays a distinct expression pattern that closely correlates with RA activity and with the dynamics of RA-signaling. ALDH1A2 is responsible in the mouse for almost all RA production during early embryogenesis, i.e., until~E8.5 [13]. During gastrulation, ALDH1A2 is expressed mainly along the primitive streak and in mesodermal cells in the posterior end of the embryo [26]. Later, ALDH1A2 is expressed in the somatic and lateral mesoderm, posterior heart tube, and rostral forebrain-and subsequently in prospective cervical and trunk levels during body axis extension [26]. Thereafter, ALDH1A3 is responsible for RA synthesis in the eye and olfactory system. ALDH1A1, thought to be partly redundant with ALDH1A3, has been demonstrated to act only during eye development [27,28] Alternative pathway of RA synthesis An alternative pathway for RA synthesis (elucidated in zebrafish) involves conversion of β-carotene to retinaldehyde by a β-carotene cleaving enzyme, β-carotene 15, 15'-oxygenase 1 (BCO1) [29]. This pathway, believed to be an ancestral pathway from early chordates, is found mainly in marine fish in which retinaldehyde and Fig. 1 Retinoic acid (RA)-signaling pathway. Extracellular binding and import of retinol, retinol conversion, oxidation of retinaldehyde to ATRA and/or 9-cis-RA, binding to RA receptors (RAR, RXR), degradation of RA to metabolites, and activation/repression of retinoic acid response elements (RARE). This diagram shows the activation (Arrow) of a target gene (Red box). Lightning bolts indicate genes with associated human eye development diseases. Abbreviations: RBP4, retinol-binding protein 4; STRA6, stimulated by retinoic acid 6; RDH5, retinol dehydrogenase 5; RDH10, retinol dehydrogenase 10; ADH1, alcohol dehydrogenase 1; ADH7, alcohol dehydrogenase 7; CRBP, cellular retinol-binding protein; ALDH1A1, aldehyde dehydrogenase 1 family member A1; ALDH1A2, aldehyde dehydrogenase 1 family member A2; ALDH1A3, aldehyde dehydrogenase 1 family member A3; CYP26A1, cytochrome P450 family 26 subfamily A member 1; CYP26B1, cytochrome P450 family 26 subfamily B member 1; CYP26C1, cytochrome P450 family 26 subfamily C member 1; ATRA, all-trans-retinoic acid ; 9-cis-RA; RAR, retinoic acid receptor ; RXR, Retinoic X Receptor; RARE, retinoic acid response elements. carotenoids stored in the egg yolk are the main source of retinoids during development [29]. The mouse and human homolog, β-carotene 15,15'-oxygenase 1 (encoded by Bco1; BCO1, previously known as BCDO1), is expressed in retinal pigment epithelium (RPE)-as well as in kidney, intestine, liver, brain, stomach, and testis [30][31][32]. Its primary function is to generate retinaldehyde in photoreceptor cells and to supplement retinoid pools in other tissues [33].
In addition, a second β-carotene cleavage enzyme expressed in rabbits, β-carotene 9',10'-dioxygenase (encoded by the BCO2 gene), catalyzes the cleavage of β-carotene into β-apocarotenoic acid, which can be transformed into RA without any involvement of ALDHs [34]. Lastly, cell culture studies have revealed that CYP1B1, a member of the cytochrome P450 family, can catalyze conversion of retinol to retinaldehyde and RA. It remains to be seen if this enzyme meaningfully contributes to RA synthesis in mammals [35].
Although it was originally shown that the RA CYP26mediated polar derivative-4-oxo-RA-can interfere with embryonic development when delivered exogenously by binding to and activating RARs [41], more recent in vivo data suggest that CYP26-mediated catabolism is required for embryonic development because its removal of RA prevents inappropriate signaling in specific tissues [42]. The CYP26 enzymes display an expression pattern which matches that of the ALDHs during embryogenesis; their targeted disruption causes teratogenic effects similar to those seen in RA toxicity. In mice, Cyp26a1 and Cyp26c1 are the first genes to be expressed in the rostral-most embryonic epiblast, whereas Cyp26b1 is expressed in tail bud tissues and in the distal limb bud mesenchyme. Later in development, these enzymes display differential expression patterns in various developing organs, such as retina, dental epithelium, and inner ear [reviewed in [25]].

RA gene activation or repression
RA acts as an agonist of two nuclear receptor families that bind DNA and directly regulate transcription (Fig. 1). These families are (i) the RA receptors, i.e., retinoic acid receptor alpha (RARA), retinoic acid receptor beta (RARB) and retinoic acid receptor gamma (RARG), and (ii) the retinoid X receptors, i.e., retinoid X receptor alpha (RXRA), retinoid X receptor beta (RXRB), and retinoid X receptor gamma (RXRG) [reviewed in [43]]. The RARs are highly conserved in vertebrates and are primarily activated by all-trans-RA (ATRA). By contrast, the RXRs are activated by 9-cis-RA, a stereoisomer of ATRA that is detected only when vitamin A is in excess. RXRs are thought to act as heterodimeric scaffolding proteins that facilitate binding of the RAR-RXR complex to DNA-complex demonstrates greater affinity for DNA than the RAR or RXR homodimers [44][45][46]. RARA, RXRA, and RXRB are widely expressed in tissues, suggesting that most tissues are potential targets of RA [reviewed by [47]]. Mouse knockout studies involving the RAR and RXR families have shown developmental abnormalities when two or more receptors are inactivated with the exception of RXRA-null mice which die in utero (vide infra), suggesting a degree of functional redundancy [25].
The retinoic acid pathway regulates eye development RA-signaling in mammalian eye development has been previously reviewed [54]. As such, we will focus on ocular developmental diseases associated with perturbed RAsignaling.
The process of eye development is largely conserved among chordates-including zebrafish, mice, and humans [55,56]. Mouse eye development begins at E8.0, at which time the optic vesicle forms on the cephalic neural folds [57]. The optic vesicle then begins to migrate towards the surface ectoderm until, at E9.0, the two ectodermal layers come into contact and begin to thicken. This contact initiates activation of a cascade of transcription factors (e.g., SIX3 and PAX6) [reviewed in [58]] and signaling pathways [e.g., BMP and RA (reviewed in [59]].
The optic vesicle then invaginates into the optic cup, and the surface ectoderm subsequently invaginates to form the lens placode. As the lens placode continues to invaginate, asymmetric cell growth then leads to formation of the lens pit, with the ultimate formation of the lens vesicle by E11. Epithelial cells located at the anterior portion of the lens vesicle maintain their epithelial identity and proliferative nature, whereas epithelial cells at the posterior lens vesicle differentiate into fiber cells and ultimately become primary lens fiber cells. The inner layer of the optic vesicle then becomes the neuroretina, while the outer layer becomes the RPE [60].
From mouse neuroretina, several neuronal subtypes (i.e., retinal ganglion cells, amacrine cells, horizontal cells, bipolar cells, photoreceptor cells) and Müller cells begin forming at E11. Corneal development begins when the lens stalk connecting the lens vesicle to the surface ectoderm is severed. The resulting space is rapidly filled by invading cells from perinuclear mesenchyme. Mesenchymal cells nearest the lens vesicle then form the corneal endothelium, while cells farthest from the lens vesicle form the corneal epithelium. Cells located between these two layers form the corneal stroma from which corneal keratocytes differentiate.
Corneal development is maintained by a constant influx of cells from the periocular mesenchyme (POM). Lens formation continues, with secondary fiber differentiation in the mouse beginning at the lens equator at E13.5-E14.5 (secondary fiber cell differentiation occurs throughout adulthood). Anterior segment development is completed by the anterior edge of the optic cup (which forms the epithelium of the iris and ciliary body), and migrating cells from the POM form stroma of the iris and ciliary body. Lastly, the trabecular meshwork is formed from migrating mesenchymal cells. Eye morphogenesis is largely conserved in mouse, zebrafish, and humans, but the process in zebrafish occurs in a much shorter time frame [56,61].
Studies in animal models have revealed a requirement for RA-signaling in normal eye development. Beginning in the mid-twentieth century, research highlighted the importance of dietary vitamin A in maintaining rodent eye development [3,62]. Rats born to mothers maintained on a vitamin A-deficient diet displayed a multitude of ocular defects-including retina infolding, coloboma, microphthalmia, and anophthalmia (a syndrome ultimately termed vitamin A deficiency)-that could be rescued by vitamin A supplementation during embryonic development [3,62]. However, the time of supplementation was critical, in that supplementation before E13.0 could rescue the ocular phenotypes whereas supplementation after E13.0 could not completely rescue eye development [3]. These studies demonstrated that vitamin A (and ultimately RA-signaling) is required for specific events in eye development, i.e., optic cup formation, anterior segment formation.
As noted, RA-signaling is modulated by several enzymes and dependent on RARs and RXRs (Fig. 1). Therefore, an alteration in any one of these proteins may perturb RA-signaling and affect normal eye development. Animal models used to investigate the RA-signaling pathway will be discussed in the order that the proteins appear within the pathway-starting with RBP4 and finishing with RARs and RXRs (Fig. 1).
In zebrafish, decreased stra6 (e.g., by morpholino knockdown) causes reduced eye size, despite the formation of all retinal layers [63]. In mouse, RDH10 was identified as essential for normal eye development [24]. RDH10deficient mice lack the cornea and ventral half of the retina and exhibit hypoplastic lenses. RDH10-depleted zebrafish display a mild RA loss of function phenotype [64], perhaps due to their ability to produce retinaldehyde through bcox [29]. A morpholino-mediated inhibition of bcox results in microphthalmic zebrafish with a diminished size of the ventral prospective retina tissue [29].
In mice, ALDHs are differentially required at various stages of eye development. ALDH1A2 is only expressed in the murine eye between E8.5 and E9.5 and is required for optic cup formation (Fig. 2a) [65]. In contrast, ALDH1A1 and ALDH1A3 are respectively expressed in dorsal and ventral retina from E10.5 onwards [27] (Fig. 2b). Aldh1a1-null mice exhibit no developmental ocular phenotype, likely due to compensation in RA-signaling by ALDH1A3 [27,28]. Aldh1a3-null mice display developmental ocular phenotypes-resulting as ventral rotation of the lens, persistence of the primary vitreous, and thickening of the ventral POM [27,66]. Aldh1a1/Aldh1a3-null mice display all of the same phenotypes as Aldh1a3-null mice, but with greater severity; this would suggest that some of the loss of RA-signaling induced by the genetic ablation of Aldh1a3 is compensated by Aldh1a1 [27]. RA influences mammalian ocular development in a paracrine fashion; RA produced in the retina by ALDH1A1 and ALDH1A3 is secreted and acts on cells in the POM where it regulates expression of genes important for apoptosis and corneal morphogenesis and cell specification-Eya2 and Pitx2, respectively (Fig. 2b) [27,67,68].
Disruption in RA-signaling permits overgrowth of POM cells, which adversely affects normal anterior segment development [27,28]. Ectopic lens expression of CRABP1 results in lenses with impaired secondary fiber cell differentiation (i.e., failure to lose nuclei) and a flattening of the anterior side of the fiber cells [69]. ATRA binds to and activates the RXR/RAR complex, which enables activation or repression of RARES (Fig. 1). Compound Rar gene deletions (for example, the deletion of both Rara and Rarb) result in aberrant ocular developmental phenotypes-including microphthalmia, coloboma, lens abnormalities, and retinal dysplasia and degeneration [70,71].
Compound Rxra, Rxra/Rarg, and Rxra/Rara null mice all show ocular developmental abnormalities-including ventral rotation of the lens, thicker corneas, shorter ventral retinae, and coloboma [72]. Involvement of RA-signaling in maintenance of POM cellular proliferation was confirmed by the conditional deletion of Rara, Rarb, and Rarg in neural crest cells; these knockout animals have impaired ocular development phenotypes similar to the previously described ALDH-deficient mice [68]. Collectively, these animal models have provided compelling evidence in support of an important role for RAsignaling in ocular development.
Mutations in genes involved in RA-signaling in humans are associated with developmental diseases-including the ocular developmental diseases, such as microphthalmia, anophthalmia, and coloboma (collectively called MAC disease), and Mathew-Wood Syndrome (Fig. 1). Linkage analysis and whole-exome sequencing have identified mutations in RBP4 in patients with MAC disease [73,74]. Dominant-negative mutations in RBP4 increase RBP4 affinity for STRA6; this nonproductive occupation of STRA6 hinders delivery of vitamin A to the fetus. Maternal inheritance of RBP4 mutations and a lack of maternal dietary retinoids predispose the fetus to MAC disease [73]. Mutations in STRA6 are associated with both syndromal (Mathew-Wood Syndrome) and non-syndromal MAC disease [75][76][77][78]. A double-nucleotide polymorphism that causes a nonsynonymous change from glycine to lysine in a highly conserved region of the STRA6 protein was identified in MAC patients; this mutation almost completely abolishes cellular uptake of vitamin A [75].
Homozygous nonsense mutations, missense, and splice-site mutations in ALDH1A3 are associated with Fig. 2 The retinoic acid (RA)-signaling pathway and eye development. Key events in mouse eye development that are regulated by RA-signaling are shown. a Between E8.5 and E9.5, ALDH1A2 is expressed in the optic vesicle and is required for optic cup formation [65]. b Starting at E10.5, RA produced by ALDH1A1 and ALDH1A3 within the developing retina diffuses (shown by green arrows) into periocular mesenchyme (POM) where it activates RARBs/RARGs on POM cells to induce cell apoptosis and thus control the migration of POM cells to the anterior eye. This diagram represents conditions at E11.5 [inspired by [27]. Abbreviations: ALDH1A1, aldehyde dehydrogenase 1 family member A1; ALDH1A2, aldehyde dehydrogenase 1 family member A2; ALDH1A3, aldehyde dehydrogenase 1 family member A3; E, embryonic day microphthalmia [79][80][81]. Co-transfection of wild-type and mutated human ALDH1A3 (c.265C>T and c.1477G>C) revealed that the mutated ALDH1A3 protein is likely unstable and subject to proteasomal degradation [79]. Mutations in RARB have been identified in patients with syndromic MAC [i.e., pulmonary hypoplasia/agenesis, diaphragmatic hernia/eventration, anophthalmia/microphthalmia, and cardiac defect (PDAC)] [82,83]. Evaluation of these mutations have indicated that both gain-of-function and dominant-negative mutations within RARB can cause PDAC syndrome [82]. A recent report identified a de novo mutation in RARA in a coloboma patient which is hypothesized to impair the interaction between RA and RARA [84].
It is clear that RA-signaling is similarly important for zebrafish, mouse, and human eye development. Given that it is unethical to investigate the role of RA-signaling in human eye development in a manner similar to that in animal models, it is currently unknown exactly how RA-signaling might impact human eye development. However, the conserved nature of RA-signaling and eye development across chordates suggests that RAsignaling is also very likely to act in a paracrine fashion to regulate eye development in humans.

Deleterious clinical variations in the RA pathway
The ExAC database (comprising whole-exome sequencing of more than 60,000 individuals) was published in 2016 [85]; it was recently expanded to include~140,000 individuals [16]. Using this resource, it is now possible to gain insights into the necessity of various components of the RA pathway in humans and to explore the existence of nodes in the pathway that are of potential importance.
A gene with a high pLI score [85] would suggest that individuals who inherit loss-of-function (LOF) mutations in that gene will inherit a survival disadvantage. Analysis of pLI scores of genes involved in the RA-signaling pathway (Table 1) reveals that certain genes in this pathway are crucial for human embryogenesis and life-data that are consistent with mouse studies ( Table 2). The RAsignaling pathway is essential for life, with 30% of RA-signaling pathway genes being categorized as LOF-intolerant. This is in striking contrast to 17% of all known genes being categorized as LOF-intolerant [85]. Several of the LOF-intolerant RA-signaling pathway genes have no known associated human disease, i.e., RDH10, RXRA, RXRB (Table 1); this would be predicted given the severe embryonic lethality observed in transgenic mouse models in which these three genes have been knocked out (Table 2).
pLI predictions and animal models are in agreement for one gene in particular-ALDH1A2. Humans are LOF-tolerant (pLI = 0.36) ( Table 1) and Aldh1a2 heterozygous null mice are viable. This highlights an important feature of pLI scores; they predict the probability of haploinsufficiency intolerance [158]. However, it is important to note that Aldh1a2-null mice experience embryonic lethality by E10.5 (Table 2) [111].
Despite the high level of conservation in the RA-signaling pathway between humans and mice, discrepancies in the extent of indispensability of RA-signaling pathway genes exist. ALDH1A1 is intolerant of LOF mutations (pLI = 0.95) in humans (Table 1) whereas Aldh1a1 is dispensible in mice [27]. This may be related to loss of Aldh1a1 being compensated in mice by Aldh1a7-which is an Aldh1a1 gene duplication found in rodents but not in humans [159].
In mice, Cyp26b1 deletion is lethal immediately after birth [123,124], whereas humans with CYP26B1 mutations can live to adulthood [96]. Cyp26a1-null mice experience embryonic lethality (Table 2), while human CYP26A1 is LOF-tolerant (pLI = 0) ( Table 1). Differences between mouse [160] and human [161] CYP26A1 tissue expression may explain the differential requirements for life. For example, human MAP2-positive neurons in the human dentate gyrus express CYP26A1, whereas rat and mouse MAP2-positive neurons do not [162]. Further, human MAP2-positive neurons express ALDH1A2 along with CYP26A1, suggesting that RA acts in an autocrine fashion in these cells, as opposed to the paracrine fashion found in rodents. This may explain the differences in the requirement for life. Differences between mouse and human underscores the need for caution when generalizing the requirement for life of RA-signaling pathway genes across chordates. Despite this, mice still hold great utility as an experimental model when investigating the minutiae of the RA-signaling pathway.
Differences between animal models and humans can be further explained by incomplete penetrance in human-possibly due to differences in mutation type, variations in gene expression, epigenetic changes, age, sex, or copy number variations [163]. Often, experimental models used for investigation of the RA-signaling pathway rely on transgenic mice in which a pathway gene is completely ablated. This situation may not be representative of humans in whom gene mutations commonly result in lowered gene activity rather than zero expression. In addition, important differences during development in the patterns of expression of many genes and their pathways exist between mouse and human [164][165][166]. As was highlighted above, differences in the timing and/or location of RA-signaling pathway genes result in vast differences in this critical pathway-e.g., Table 1 Genes involved in the retinoic acid synthesis, degradation, and signaling in humans  Increased energy dissipation, insulin resistance, diet-induced obesity resistance [108], significantly reduced ability to convert retinol to retinoic acid in the liver [109] Aldh1a2 RALDH2 107928 Devoid of retinoic acid, die by E10.5 with impaired hindbrain development, failure to turn, lack of limb buds, heart abnormalities, reduced otocysts and a truncated frontonasal region [110][111][112][113][114] Aldh1a3 RALDH3 1861722 Neonatal death [115], persistent hyperplastic primary vitreous, thick neural retina and no vitreum [28], choanal atresia, ethmoturbinal hypoplasia, ventral lens rotation, short ventral retina, and no Harderian gland [66] Aldh1b1 ALDHX 1919785 Increased fasting circulating glucose levels and decreased blood acetaldehyde clearance [116], defects in beta cell development and functionality, glucose intolerance, agedependent hyperglycemia, and insulin resistance [117] Aldh8a1 ALDH12 or RALDH4 2653900 None Found Cyp1b1 88590 Protected from the acute bone marrow cytotoxic and preleukemic effects of DMBA [118], show a decreased incidence of DMBA-induced lymphomas [119], ocular drainage structure abnormalities (~POAG) [120] Cyp26a1 1096359 Mid-late gestation lethal and exhibit spina bifida, caudal agenesis, and abnormalities of the kidneys, urogenital tract, hindgut, cervical vertebrae, and rostral hindbrain [121,122], rescued by partial Aldh1a2 deletion [42] Cyp26b1 2176159 Lethal immediately after birth exhibiting respiratory distress [123], limb morphogenesis and proximal-distal patterning is disrupted in homozygous null fetuses [123,124], abnormal spermatogenesis/oogenesis [125], arrested hair follicle development [126] Cyp26c1 2679699 Viable, exhibit normal CNS development with no apparent anatomical defects [127] Crabp1 CRABP-I 88490 Phenotypically normal and fertile [128] Crabp2 CRABP-II Postaxial polydactyly [129] Fabp5 101790 Impaired skin barrier function [130], resistance to dietinduced obesity (decreased adipose tissue and improved glucose tolerance) [131], and impaired cognitive function [132,133] Rara NR1B1 97856 High neonatal mortality due to maternal cannibalization, failure to thrive, and excess mortality during the postnatal period, male survivors exhibit testicular degeneration [134][135][136] Rarb NR1B2 97857 Reduced growth [137], locomotion abnormalities [138] Rarg NR1B3 97858 Stunted growth, homeotic transformations of the rostral axial skeleton and tracheal cartilage, Harderian gland agenesis, high postnatal mortality, and male sterility [135,139], splenomegaly, and myeloproliferative disease, abnormal granulocytes [140] Rbp1 CRBPI 97876 Increased adiposity, increased PPAR-gamma target gene expression [141], increased Rpb2 and Crabp2, elevated pancreatic RA [142], increased susceptibility to a diet deficient in vitamin A [143] Rbp2 CRBPII 97877 Pups of homozygous dams fed a marginal retinol diet show increased neonatal lethality due to inadequate retinal transport to the fetus, abnormal retinol level and vitamin absorption [144] autocrine vs. paracrine signaling, embryonic or postnatal lethality, and tissue-specific expression (or lack thereof). All of these can contribute to the discrepancies between mouse models for the RA-signaling pathway genes, observed human diseases, and the pLI scores.
Future studies of the retinoic acid signaling pathway By means of analysis of genetic intolerance, we can pinpoint certain members of the RA-signaling pathway that are likely to be essential for human life. Clearly, it is important that these members need to be better understood. This can be achieved by generating hypomorphic mutations in mice, i.e., protein function is diminished rather than ablated. Hypomorphic mutations in mice can be studied via the introduction of single-nucleotide variants (SNVs) using CRISPR/Cas9 technology [reviewed in [167]]. This approach was recently used to study hypomorphic mutations in a humanized CYP3A5*1 mouse model [168]. These humanized mice with hypomorphic mutations are likely to be better models of human diseases associated with altered RA-signaling pathways. While CRISPR/Cas9 can be effectively used for modeling human diseases in mice, such studies can be prohibitively inefficient in that they use advanced techniques, need specialized equipment, and require at least 3 months to generate knockin/knockout mice [169]. Zebrafish represent an alternative in which high-throughput screening can be used to investigate mutations in genes of the RA-signaling pathway that have been identified in humans [ [170] and reviewed in [171]]. Zebrafish are more efficient than mice for such CRISPR/Cas9 experimental approaches for several reasons: they have a shorter generation time, produce more offspring, and are less expensive to maintain [172][173][174][175]. CRISPR/Cas9 was recently used in zebrafish to generate a humanized model of renal agenesis in which GREB1 like retinoic acid receptor coactivator (GREB1L) was identified as a coactivator of RAR genes [176]. It is expected that similar approaches will be used in future studies to manipulate the RA-signaling pathway and thereby enhance our understanding of RA-signaling in human physiology and pathophysiology.

Conclusions
For centuries, the importance of dietary vitamin A to human health has been known. Ancient civilizations unknowingly used homeopathic remedies in which vitamin A was the main active ingredient. It was not until the turn of the twentieth century that our more nuanced understanding of the role for vitamin A in human health began to take shape. Pioneering animal studies determined that vitamin A was critical for embryogenesis, eye development, and identified retinoids as derivatives of vitamin A. Decades later, our understanding of the RAsignaling pathway has grown significantly and now includes a more comprehensive knowledge of retinol cellular uptake and oxidation, RA catabolism, nuclear receptor (RAR/RXR) activation, and nuclear receptor gene targets-and the importance of the RA-signaling pathway for eye development. By leveraging information gained from large-scale human whole-exome sequencing efforts (e.g., ExAC and gnomAD), our understanding about the importance of the RA-signaling pathway to human health is improving. This was underscored by the high number of genes within this pathway with large pLI scores. While transgenic mouse models have provided valuable insights into the details of the RA pathway, discrepancies between the human and mouse data Rdh5 1201412 Impaired dark adaptation and at high bleaching levels, large increase in 11-cis-retinyl ester concentration [147] Rdh10 1924238 Mid-gestational lethality, reduced RA-signaling and abnormal limb, craniofacial, somite and cardiac morphology including microphthalmia [24] and dorsal pancreas agenesis [148] Rxra NR2B1 98214 Multiple organ defects and die of cardiac failure by E14.5, eye defects (retinal abnormalities, late corneal opacity), placental defects [149][150][151][152][153] .
Rxrb NR2B2 98215 Partial embryonic and perinatal lethality, and surviving adult males are sterile due to defects in spermatogenesis [154] Rxrg NR2B3 98216 Neuron reduction in striatum, premature death and altered responses to the administration of dopamine antagonists [155] Stra6 107742 Seven-fold reduction in total ocular retinoids, photoreceptor anomalies, abnormal RPE, sclera, and choroid [20] underscore the need for care when generalizing results from animal studies to humans. Animal models will continue to enhance our understanding of the RA-signaling pathway under physiological and pathophysiological conditions. Exciting new models and techniques (e.g., Zebrafish, CRISPR/Cas9, hypomorphic mutations) will allow a more nuanced examination of this pathway. These will allow the pathophysiological consequences of individual human mutations in the RA-signaling pathway to be explored. In summary, the RA-signaling pathway play a critical role in embryogenesis, eye development, and are required for life. We should anticipate fascinating new insights into this pathway in the coming years.

Acknowledgements
We thank the Yale Printing and Publishing Service for their assistance with Fig. 2, and also appreciate our colleagues for careful reading of this manuscript and constructive comments.
Authors' contributions BT participated in writing and editing the manuscript and generating Tables 1 and 2 and Figs. 1 and 2. NK participated in writing and editing the manuscript and providing insightful feedback. NA participated in writing the manuscript and generating Tables 1 and 2. DCT and DWN participated in writing and editing the manuscript, and providing insightful feedback. VV participated in conceiving the idea for the manuscript and in drafting, editing, and finalizing the manuscript.

Funding
This work was supported in parts by National Institutes of Health Grants AA021724, AA022057, EY017963, and EY022312. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.