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. 2017 Mar 31;26(13):2386–2397. doi: 10.1093/hmg/ddx127

Loss of dynein-2 intermediate chain Wdr34 results in defects in retrograde ciliary protein trafficking and Hedgehog signaling in the mouse

Chuanqing Wu 1,2, Jia Li 2,3, Andrew Peterson 4, Kaixiong Tao 1,*, Baolin Wang 2,5,*
PMCID: PMC6075199  PMID: 28379358

Abstract

The Wdr34 gene encodes an intermediate chain of cytoplasmic dynein 2, the motor for retrograde intraflagellar transport (IFT) in primary cilia. Although mutations in human WDR34 have recently been reported, the association of WDR34 function with Hedgehog (Hh) signaling has not been established, and actual cilia defects in the WDR34 mutant cells have also not been completely characterized. In the present study, we show that Wdr34 mutant mice die in midgestation and exhibit open brain and polydactyly phenotypes. Several Hh-dependent ventral neural cell types are not specified in the mutant neural tube. The expression of the direct Hh targets, Gli1 and Patched 1, is inhibited, while the expression of limb patterning genes that are normally inhibited by the Gli3 repressor is anteriorly expanded in mutant limbs. Comparison of cilia phenotype and function among wild type, Dnchc2 (dynein 2 heavy chain), and Wdr34 mutant cells demonstrates that cilia in both Dnchc2 and Wdr34 mutant cells are stumpy. Several ciliary proteins examined abnormally accumulate in the cilia of both mutant cells. Consistent with its function, overexpressed Wdr34 is occasionally localized to cilia, and Wdr34 is required for the ciliary localization of dynein 2 light intermediate chain Lic3. More interestingly, we show that both Dnchc2 and Wdr34 act between Smo and Gli2/Gli3 in the Hh pathway. Therefore, like Dnchc2, Wdr34 is required for ciliogenesis, retrograde ciliary protein trafficking, and the regulation of Gli2/Gli3 activators and repressors. Furthermore, both Wdr34 and Dnchc2 promote microtubule growth, a novel dynein 2 function in a non-cilia structure.

Introduction

The family of secreted Hedgehog (Hh) molecules plays an important role in the development of the central nervous system, limb, and many other structures in vertebrates (1). Hh signaling is initiated by the binding of Hh ligand to its cell surface receptor, Patched (Ptch), a twelve-pass membrane protein (2,3). This binding prevents Ptch from inhibiting Smoothened (Smo), a G-protein coupled receptor, allowing the Hh pathway to be activated. Smo then transmits signals to activate the downstream Gli transcription factors. Of three Gli family members, Gli2 and Gli3 primarily mediate Hh signaling. In the absence of Hh signaling, Gli2 and Gli3 full-length proteins (Gli2FL and Gli3FL) are proteolytically processed to generate C-terminally truncated transcriptional repressors (Gli2Rep and Gli3Rep) (4,5). Hh signaling inhibits the proteolytical processing and converts the latent full-length proteins into activators, which subsequently activate the downstream transcriptional targets. Interestingly, one of these targets is Gli1, which serves to further enforce the Hh pathway activation, while Ptch1 is another target to establish a negative feedback regulation of the pathway.

It is generally accepted that vertebrate Hh signaling occurs in the primary cilium, a solitary microtubule-based organelle that protrudes from the cell surface and functions in transducing extracellular signals (6). Consistent with this, the Hh pathway core components, including Ptch1, Smo, Gli2 and Gli3 proteins, Sufu, and Kif7, are localized to the cilia (7–11). Ciliary defects cause a diverse array of developmental abnormalities, such as polydactyly, central nervous system defects, polycystic kidney disease, respiratory and visual disorders, hydrocephalus, obesity, and mental retardation, which are collectively termed ‘ciliopathies’ (12).

Primary cilia originate from the mother centrioles, which serve as a basal body and a docking station for ciliary microtubules and intraflagellar transport (IFT) machinery during primary cilia elongation. IFT machinery is also essential for the maintenance of primary cilia. There are two IFT protein complexes, IFT-A and IFT-B. IFT-B, together with a kinesin 2 motor, is responsible for anterograde ciliary trafficking of protein complexes and vesicles, while IFT-A, together with a cytoplasmic dynein 2 motor, is responsible for retrograde ciliary protein and vesicle trafficking (13,14). There are several kinesin 2 motors that power anterograde ciliary trafficking (15). In contrast, cytoplasmic dynein 2 is currently the only known retrograde IFT motor in cilia. Dynein 2 in metazoans consists of a dimer of heavy chain subunits (encoded by Dnchc2, also known as Dync2h1, Dhc1B, or Dhc2), which associate with intermediate chains, light intermediate chains, and light chains (16–18). Mutations in mouse Dnchc2 results in stumpy cilia and Hh signaling defect (19,20), and mutations of human DYNC2H1 gene also cause short-rib polydactyly syndrome type III (21). Similarly, mutations in human WDR34 gene, which is recently shown to encode a dynein 2 intermediate chain (16), also result in short-rib polydactyly syndrome type III (22,23). Human WDR34 mutant fibroblasts exhibit short and bulbous cilia. However, neither has it been established whether WDR34 mutations affect Hh signaling, nor has the mouse Wdr34 mutant phenotypes been reported.

In this study, we show that loss of Wdr34 in mice results in stumpy cilia and abnormal accumulation of ciliary proteins in cilia. Wdr34 mutant mice exhibit open brain and polydactyly phenotypes. Nevertheless, although Gli2FL and Gli3FL levels are increased in the mutant, they are not activated, as the expression of Hh targets that is dependent on activated Gli2FL and Gli3FL is inhibited. On the other hand, the expression of the limb patterning genes that are normally inhibited by Gli3Rep are anteriorly expanded in the mutant limb bud, suggesting that Gli3Rep function is also attenuated. We also show that Wdr34 acts between Smo and Gli2/Gli3 in the Hh pathway. Thus, our study links Wdr34 function to Hh signaling, ciliogenesis, and retrograde ciliary trafficking.

Results

Loss of Wdr34 impairs Hh signaling in mice

To determine the role of Wdr34 in vivo, we used the targeted gene homologous recombination approach to mutate the gene by deleting exons 3–8 in mice (Fig. 1A and B). The deletion of the exons, which cover most of the coding region, is expected to cause an open reading frame shift, thus most likely resulting in a null allele. The vast majority of mice homozygous for the Wdr34 mutation died in midgestation varying from embryonic day 10.5 to 12.5 (E10.5 –E12.5), occasionally surviving until E16.5. The mutant embryos exhibited open brain, spinal bifida, microphthalmia, and polydactyly (Fig. 1C and D).

Figure 1.

Figure 1

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Inactivation of Wdr34 results in open brain, polydactyly, and microphthalmia in mice. (A) The gene targeting strategy used to create a mouse Wdr34 mutant allele. Open rectangles are referred to as exons and lines as introns. Probe and BamHI restriction sites used for Southern blot are shown. Triangle, loxP site; Neo, neomycin; DTA, diphtheria toxin A. (B) Southern blot of representative mutant and wild type (wt) ES cell clones. (C) The morphology of wt and Wdr34 mutant embryos at different developmental stages. Note the delayed development, open brain (pointed by arrowhead), and microphthalmia (indicated by arrow) of mutant embryos. (D) Limb phenotypes of E16.5 and E12.5 wt and mutant embryos. Note that E12.5 mutant limb paddle is wider than that of wt, indicating polydactyly.

Given the role of Hh signaling in the central nervous system, eye, and limb development, we next characterized the neural tube phenotype of the mutant in more detail. The generation of all five types of ventral neural progenitors in the developing neural tube is induced by different concentrations of Sonic Hedgehog (Shh) that is expressed in the notochord and the floor plate, while the dorsal neural progenitor specification is inhibited by Shh signaling. These different types of neural progenitors can be marked by the expression of specific transcription factors (24). We thus examined the expression of several neural tube markers in Wdr34 wild type and mutant neural tube of E10.5 embryos by immunofluorescence. In wild type neural tube, Foxa2, Nkx2.2, and Hb9 and Isl1 label the floor plate, V3 progenitors, and motor neurons, respectively. However, all these ventral markers were absent in the Wdr34 mutant neural tube. In contrast, Pax6 expression, which is normally ventrally restricted by the high Shh signaling, expanded throughout the entire ventral neural tube. Similarly, dorsal marker Pax7 expression domain was also slightly ventrally extended. On the other hand, loss of Wdr34 does not affect Shh expression in the notochord, indicating that a lack of the ventral neural tube markers’ expression was due to defective Hh signal transduction but not Shh expression (Fig. 2A).

Figure 2.

Figure 2

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Inactivation of Wdr34 disrupts Shh-dependent neural patterning in the neural tube and inhibits Hh signaling. (A) Ventral neural progenitors are not specified in the Wdr34 mutant neural tube. E10.5 neural tube sections were immunostained for the indicated protein markers (red) and counterstained with DAPI for nuclei (blue). Note that none of the ventral markers was detected in the mutant, while the expression of dorsal markers (Pax6 and Pax7) was ventrally expanded. The two equal long lines are used to compare the length of ventral region lacking Pax7 expression in wt and the mutant, which is slightly shorter in the mutant. Arrowhead and arrow indicate Shh expression in the floor plate and notochord, respectively. (B) Ptch1-lacZ expression is undetectable in the mutant. E10.5 embryos with indicated genotypes were subject to lacZ staining. Transverse neural tube sections are shown to the right.

To determine whether inactivation of Wdr34 directly affects Hh signaling, we examined the expression of Ptch1, a transcriptional target of Hh signaling. To this end, the Wdr34 mutant allele was crossed into Ptch1lacZki heterozygous mutant, in which the lacZ gene is inserted in the Ptch1 locus so that lacZ expression is an indicator of Ptch1 expression (25). As predicated, lacZ was expressed in brain, spinal cord, and limb buds of wild type E10.5 embryos, but not in those of Wdr34 mutants (Fig. 2B). Similarly, in situ hybridization results showed that the expression of both Ptch1 and Gli1, another Hh target, was detected only in wild type but not the mutant limb buds, though Shh expression was unaffected (Fig. 3).

Figure 3.

Figure 3

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Altered expression of limb patterning genes in Wdr34 mutant limb buds. Whole mount in situ hybridization of E10.5 limb buds shows the expression of limb patterning genes. Note that Ptch1 and Gli1 expression is not detected, while Shh is normally expressed in the mutant. The expression of the genes that are normally inhibited by Gli3 repressor is anteriorly expanded (Gremlin, Hand2, Hoxd11, Hoxd13, Fgf4, and Fgf8). Note that although the higher expression domain of Hand2 in the mutant limb is smaller than that in WT limb, the weak expression is anteriorly expanded. Arrows indicate the anterior end of Fgf4 and Fgf8 expression.

To determine the molecular basis of the polydactyly phenotype of the Wdr34 mutant, we examined expression of several limb-patterning genes that are regulated by Hh signaling. Gremlin, Hand2, Hoxd11, and Hoxd13 expression was normally restricted to the posterior region of developing limbs by Gli3Rep (26–29). However, the expression of all these genes was anteriorly expanded in the mutant limbs. Similarly, the expression of both Fgf4 and Fgf8 in the apical ectodermal ridge (AER), which is also restricted by Gli3Rep (30), was anteriorly extended in the mutant limb as compared to wild type (Fig. 3).

To correlate the mutant phenotypes with Gli2 and Gli3 functions, we examined Gli2 protein levels and Gli3 processing in Wdr34 mutant embryos by immunoblotting with Gli2 and Gli3 antibodies (5,31). The results showed that levels of both Gli2FL and Gli3FL were increased in the mutant, while Gli3Rep levels were reduced (Fig. 4). Quantitative analysis showed that the Gli3FL to Gli3Rep ratio in the mutant is markedly increased as compared to that of wild type (graph in Fig. 4). Thus, Gli3 processing is reduced in the Wdr34 mutant. Since the ventral neural tube patterning is primarily dependent on Gli2/Gli3 activators, whereas limb patterning mostly relies on the Gli3Rep, these results indicate that both the Gli2/Gli3 activator and repressor functions are impaired in the Wdr34 mutant.

Figure 4.

Figure 4

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Western blots show the increase in Gli2FL and Gli3FL levels and decrease in Gli3Rep levels in E10.5 Wdr34 mutant embryos. Tubulin blot is a loading control. Quantitative analysis of the Western blot results is shown by graphs to the right.

Loss of Wdr34 and Dnchc2 results in stumpy cilia and defects in retrograde ciliary trafficking

Given that cilia are required for Hh signaling and Wdr34 is an intermediate chain of dynein 2, we next examined the cilia morphology by scanning electron microscopy (SEM). Cilia were long and thin in neuroepithelial cells in the lumen of the wild type neural tube, but short and swollen in the Wdr34 mutant (Fig. 5A). Immunofluorescent staining for cilia markers, Arl13b and acetylated tubulin, showed that cilia density in both neuroepithelia of the neural tube and limb mesenchyme of the mutant was similar to that in wild type. However, the staining in wild type limb mesenchymal cells was usually elongated, but dotted in the mutant, indicating that cilia in the mutant cells are short and stumpy (Fig. 5B). Consistent with this, cilia in the mutant MEFs were significantly shorter than those in wild type. Cilia number in the mutant MEFs was also significantly lower than that in wild type MEFs (Fig. 5C, upper and middle graphs). These results suggest that Wdr34 is required for ciliogenesis and maintenance of cilia length.

Figure 5.

Figure 5

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Stumpy cilia and aberrant accumulation of ciliary proteins in the cilia of Wdr34 mutant cells. (A) Scanning electron microscopy (SEM) of the cilia of neural epithelia in the neural tube of E9.5 embryos. Arrows indicate cilia, and the enlargement of framed cilia is shown to the right. (B) The neural tube and limb mesenchyme of E10.5 embryos and (C) MEFs were immunostained for the indicated proteins, acetylated tubulin (ac-Tub), and nuclei (DAPI, blue). For Smo ciliary localization, cells were treated with ShhN conditioned medium 24h prior to immunostaining. Quantitative analysis of the results in C is shown by graphs to the right. Two-tailed Student t-test shows that all p-values are equal to or less than 0.0005 (upper graph), 0.0014 (middle graph), and 0.013 (lower graph), all of which are less than 0.05, thus significant. Note that cilia in the mutant cells are short and swollen and that all ciliary proteins examined accumulate abnormally at the cilia tip of both Wdr34 and Dnchc2 mutant cells. The pointy end of ac-Tub staining is the distal end of cilia.

Given that cytoplasmic dynein 2 is the motor for retrograde IFT in cilia, it is expected that loss of Wdr34 function would lead to abnormal accumulation of ciliary proteins in cilia. To test this prediction, we examined and compared ciliary localization of several proteins in wild type, Wdr34, and Dnchc2 mutant MEFs by immunofluorescence. Both Arl13b and Smo, two ciliary membrane proteins (7,32), were distributed evenly along cilia in wild type cells. However, they accumulated at the tip of the cilia in both Wdr34 and Dnchc2 mutant cells. Similarly, Ift88, Ift57, and Ift140, which represent IFT-B and IFT-A complexes (13), respectively, were also significantly enriched in the cilia of the mutant cells (Fig. 5C, images and lower graph). It is worth noting that primary cilia shaft is made of 9 microtubule triplet in the basal body, doublet in most part of cilia, and singlet at the cilia tip (14). Thus, the staining for acetylated tubulin at the cilia tip is often pointier than that in the main part and proximal end of cilia. This becomes especially apparent in Wdr34 and Dnchc2 mutants, as cilia assembly is defective in the mutants (Fig. 5C, images with the acetylated tubulin staining in the mutant cells). Since Arl13b, Smo, Ift88, Ift57, and Ift140 all localize to the pointy end of the mutant cilia, we conclude that they accumulate at the distal tip of cilia. Thus, like Dnchc2 mutations (20,33), loss of Wdr34 disrupts retrograde ciliary protein trafficking.

Because cilia number is significantly reduced in Wdr34 mutant MEFs, we also wondered whether loss of Wrd34 would affect the centriolar localization of several centrosomal proteins. Of four proteins examined, Cep164, Odf2, Ta3, and Cep290 (34–38), none appeared to be affected, suggesting that the basal body is normally assembled in the mutant cells (Fig. 6).

Figure 6.

Figure 6

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Normal localization of centrosomal proteins in Wdr34 mutant cells. Wild type and Wdr34 mutant MEFs were coimmunostained for the indicated proteins, γ-tubulin, and nuclei (DAPI, blue).

Wdr34 acts between smo and Gli2/Gli3 in the Hh pathway

Hh stimulation leads to accumulation of Smo, Gli2, and Gli3 in cilia (7,9,11). To determine at which step in the Hh pathway that Wdr34 mutation acts, we tested whether Smo, Gli2, and Gli3 accumulated in the cilia of Wdr34 and Dnchc2 mutant cells in response to Hh stimulation. Both wild type and the mutant MEFs were incubated with ShhN conditioned medium prior to immunostaining for Smo, Gli2, or Gli3 together with acetylated tubulin. In the absence of ShhN stimulation, Smo was basically undetectable in the cilia of wild type cells we used, but detectable in about 50% and 20% of Wdr34 and Dnchc2 mutant cilia, respectively. This is consistent with the finding that loss of Dnchc2 function results in accumulation of Smo in cilia (33). Stimulation with ShhN significantly increased the percentage of Smo positive cilia in both wild type and mutant cells (Fig. 7, upper graph), indicating that loss of either Wdr34 or Dnchc2 does not impair the Smo accumulation in cilia. In contrast, Gli2 and Gli3 already accumulated in the cilia of the mutant cells without ShhN stimulation, and ShhN stimulation resulted in no further increase in Gli2 and Gli3 positive cilia number in the mutant cells, although it did in wild type cells (Fig. 7). This indicates that the loss of either Wdr34 or Dnchc2 gene function prevents Hh-dependent Gli2 and Gli3 accumulation in cilia. Thus, Wdr34 and Dnchc2 act downstream of Smo and upstream of Gli2/Gli3 in the Hh pathway. This is consistent with genetic evidence showing that ciliary genes act between Smo and Gli2 and Gli3 (6).

Figure 7.

Figure 7

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Wdr34 and Dnchc2 act between Smo and Gli2/Gli3 in the Hh pathway. WT, Wdr34, and Dnchc2 mutant MEFs were immunostained with the indicated antibodies. The cilia number with or without detectable Smo, Gli2, or Gli3 were counted. Graphs to the right show percent cilia with Smo, Gli2, or Gli3 from at least 50 cilia in three different experiments. The p-values for those bar graphs marked with asterisks are equal to or less than 0.0015 (two-tailed Student t-test). Since they are less than 0.05, the difference between –ShhN and +ShhN is significant. Note that the ciliary localization of Smo but not Gli2 or Gli3 in mutant cells responded to ShhN stimulation.

Subcellular localization of Wdr34 and its role in the ciliary localization of other subunits of the dynein 2 complex

To determine the subcellular localization of Wdr34, wild type MEFs stably expressing green fluorescent protein (GFP) tagged Wdr34, Wdr34-GFP, were selected and subjected to immunofluorescence for GFP and acetylated tubulin. Wdr34-GFP was localized to cilia in only a small number (12%, n = 35) of Wdr34-GFP expressing cells. In most cells, Wdr34-GFP appeared to be concentrated in cell cortices and filopodia-like structures. This localization is specific since GFP alone was not localized to those structures (Fig. 8A).

Figure 8.

Figure 8

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Wdr34 subcellular localization and its effect on Lic3 cilia localization. (A) Wild type MEFs stably expressing the indicated proteins were stained for GFP, acetylated tubulin (acTub), and nuclei (DAPI, blue). GFP-Wdr34 is occasionally enriched in cilia (12%, n = 35) and mostly accumulated in cell cortex and filopodia-like structure (indicated by arrows). Arrowheads indicate ciliary localization. (B) WT and Wdr34 mutant MEFs were stained for the indicated proteins. Note that Lic3 is absent in the cilia of Wdr34 mutant cells.

Wdr34 contains five WD-repeats in its C-terminus. To map the region responsible for Wdr34 ciliary localization, the N- and C-terminal regions of Wdr34 were separately fused to GFP to create GFP-Wdr34-1-220 and GFP-Wdr34-221-E expression constructs, respectively. Immunostaining of MEFs expressing these two fusion proteins showed that GFP-Wdr34-1-220 was localized to cilia in some of the GFP-WDR34-1-220 expressing cells, while GFP-WDR34-221-E was diffusely localized to the cytoplasm (Fig. 8A). These data suggest that the N-terminus controls Wdr34 ciliary localization.

The cytoplasmic dynein 2 consists of heavy chain, intermediate chain, light intermediate chain, and light chain (16). Both the heavy chain Dnchc2 and light intermediate chain Lic3 are localized to cilia (39). Given that Wdr34 is a dynein 2 intermediate chain, we were curious about whether loss of Wdr34 affected Dnchc2 and Lic3 localization to cilia. As expected, both Dnchc2 and Lic3 were localized to cilia in wild type MEFs. However, while Dnchc2 remained in the cilia of Wdr34 mutant cells, Lic3 did not (Fig. 8B). This suggests that Wdr34 is required for Lic3 ciliary localization and/or protein stability.

Delayed microtubule regrowth in Wdr34 and Dnchc2 mutant cells

Given that dynein 2 is a retrograde IFT motor along cilium axoneme, a microtubule-based structure, we were also curious about whether loss of dynein 2 affects cytoplasmic microtubule growth and dynamics. The microtubule network in both Wdr34 and Dnchc2 mutant cells was similar to that in wild type cells (Fig. 9, no treatment). To assess the microtubule growth rate, the cells were treated with nocodazole and then stained for microtubules after nocodazole removal for different period of time. Microtubules began to regrow in both wild type and mutant cells 5 min after nocodazole removal and grew slightly longer in the wild type than mutant cells in 10 min. The difference in microtubule regrowth between wild type and mutant cells became more pronounced by 20 min and remained at 60 min, albeit smaller (Fig. 9). Thus, loss of either Wdr34 or Dnchc2 results in a decrease in microtubule regrowth rate.

Figure 9.

Figure 9

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Microtubule regrowth delays in either Wdr34 or Dnchc2 mutant cells. Microtubules and nuclei were visualized with αtubulin (red) and DAPI (blue) staining, respectively. Wild type (wt), Wdr34, and Dnchc2 mutant MEFs were treated with nocodazole. Microtubule regrowth was then assessed after nocodazole removal for the indicated times. Note that nocodazole treatment destroyed microtubule organization in both wt and mutant cells (0 min) and that microtubule regrowth delayed in both mutants as compared to wt.

Discussion

Anterograde ciliary protein trafficking is powered by kinesin 2 motors, while retrograde ciliary protein trafficking is directed by cytoplasmic dynein 2 motor. A recent study showed that Wdr34 and Wdr60 are bona fide dynein 2 intermediate chains (16). Consistent with the specific role of dynein 2 in retrograde IFT trafficking in cilia, mutations in dynein heavy chain gene Dnchc2 in both human and mouse result in stumpy cilia and aberrant accumulation of ciliary proteins in cilia (19,20,33). Similarly, mutations in human WDR34 gene lead to stumpy cilia (22,23). However, cilia defects in human WDR34 mutant cells have not been completely characterized. In the present study, we show that like human Wdr34 mutations, inactivation of the mouse Wdr34 results in short and bulbous cilia. We also extend previous studies on human WDR34 by showing that all the ciliary proteins examined, including Arl13b, Smo, Gli2, Gli3, Ift88, Ift57, and Ift140, accumulate abnormally at the cilia tip of both Wdr34 and Dnchc2 mutant cells (Figs 5 and 7). Together, these data indicate that as a part of dynein 2, Wdr34 is necessary for retrograde IFT trafficking.

It is worth noting that Arl13b and Smo staining areas in the mutant cilia are usually big and loose like ‘cloud’ as compared to those of IFT protein staining (Fig. 5C). This is most likely attributed to the fact that Arl13b and Smo are localized to ciliary membrane, while IFTs are associated with the microtubule shaft in the cilia. We also noticed that cilia number is similar in the tissues examined in wild type and Wdr34 mutant embryos, but it is significantly reduced in the mutant MEFs as compared to that in wild type MEFs (Fig. 5B and C). This difference is most likely due to the difference between in vitro culture and in vivo physiological conditions that support ciliogenesis. It is also possible that since MEFs comprise only some cell types, cilia number in MEFs may not exactly represent that in embryos.

Hh signaling depends on cilia. Mutations in genes encoding kinesin motors, IFT components, dynein 2 heavy chain, and many other proteins have been shown to cause dysfunctional Hh signaling (6). Although human WDR34 mutations have been recently shown to cause stumpy cilia (22,23), the link between WDR34 and Hh signaling has not been established. Several pieces of evidence from the current study show that loss of Wdr34 gene function disrupts Hh signaling downstream of Smo, but upstream of Gli2 and Gli3. First, all Shh-dependent ventral neural markers examined (Foxa2, Nkx2.2, Hb9 and Isl1) are missing in the Wdr34 mutant neural tube. In contrast, the expression of Pax6 and Pax7, two dorsal markers inhibited by the high level of Hh signaling, is ventrally expanded (Fig. 2A). It is worth noting that unlike those in Wdr34 mutant, Hb9 and Isl1 are expressed in the neural tube of Dnchc2 mutants (20,33). This difference could be due to the difference in genetic background, potentially residual Dnchc2 mutant gene function, and/or the role in Hh signaling between Wdr34 and Dnchc2. Second, the expression of Ptch1-lacZ and Gli1, two Hh targets, is inhibited in the Wdr34 mutant neural tube and limb (Figs 2B and 3). Since the ventral neural tube patterning and the expression of the Hh targets are dependent on Gli2FL and Gli3FL activators, yet both Gli2FL and Gli3FL levels are remarkably elevated in the mutant (Fig. 4), these results indicate that Gli2FL and Gli3FL are inactive in the Wdr34 mutant. Third, Wdr34 mutant embryos exhibit open brain and neural tube and polydactyly (Fig. 1C and D), the phenotypes that are usually associated with increased Hh signaling (25,40–42). However, since both Gli2FL and Gli3FL are not activated, these mutant phenotypes are most likely due to the change in the Gli3FL to Gli3Rep ratio and/or reduced Gli3Rep function. Thus, as in Dnchc2 mutants (19,20,33), both Gli2/Gli3 activator and repressor functions are impaired in Wdr34 mutant. Lastly, loss of Wdr34 gene function does not affect the additional enrichment of Smo in cilia in response to Hh stimulation. However, it does prevent Hh-dependent Gli2 and Gli3 accumulation in cilia. This is also true for Dnchc2 mutations (Fig. 7). These results indicate that both Wdr34 and Dnchc2 act between Smo and Gli2/Gli3 in the Hh pathway.

Recent studies showed that endogenous Wdr34 is localized to the base of cilia, while GFP-tagged Wdr34 is also found in the cilia axoneme (16,23). We found GFP-Wdr34 localized to cilia only in a small fraction (12%) of MEFs stably overexpressing the fusion protein. In most of the cells, the protein appears to be enriched in cell cortices and filopodia-like structure. Similarly, GFP-Wdr34-1-220 also appears to be localized to cilia in some of GFP-Wdr34-1-220 expressing cells (Fig. 8A). This suggests that the N-terminus controls Wdr34 ciliary localization. The difference in Wdr34 localization of endogenous and overexpressed proteins from different studies could be due to different antibodies and/or cell lines used. We do not yet know whether non-ciliary localization is physiologically relevant to Wdr34 function. If it were, this would suggest that Wdr34 also functions in subcellular location(s) other than cilia. Further studies are necessary to investigate this possibility.

In addition, loss of Wdr34 appears to affect the ciliary localization of Lic3 but not Dnchc2 (Fig. 8B), suggesting that Wdr34 is required for Lic3 ciliary localization or protein stability. We could not distinguish between the two mechanisms, since we do not know whether Lic3 protein levels are reduced in Wdr34 mutant MEFs due to the sensitivity of the Lic3 antibody. Nevertheless, our finding raises the possibility that the dynein 2 entire complex, rather than individual subunits, may determine the ciliary localization and/or protein stability of individual subunits. This might also in part explain why overexpressed Wdr34 is mainly localized to the cytoplasm rather than cilia, as the amount of other subunits of the endogenous dynein 2 complex is limited.

Dynein 2 is considered to be a cilia specific motor for IFT retrograde trafficking. It is unknown whether dynein 2 plays any role in cytoplasmic microtubule network and dynamics. Our microtubule regrowth assay shows that the microtubules grow slower in Wdr34 and Dnchc2 mutant cells than that in wild type, although the microtubule network appears similar (Fig. 9). This suggests that dynein 2 promotes cytoplasmic microtubule growth. To our knowledge, this is the first piece of evidence that dynein 2 plays a role in a non-ciliary structure. Additional studies are necessary to elucidate the mechanism underlying this novel role of dynein 2.

Materials and Methods

Mouse strains and the generation of a Wdr34 mutant allele

Institutional Animal Care and Use Committee at Weill Cornell Medical College approved this research including the use of mice and mouse embryonic fibroblasts.

A BAC clone containing mouse Wdr34 genomic DNA sequences was purchased from the BACPAC Resources Center (Oakland, CA, USA) and used to create a Wdr34 targeting construct. The construct was engineered by replacing exons 3–8 of the Wdr34 gene with the neomycin cassette flanked by loxP sites (Fig. 1A). The linearized construct was electroporated into W4 ES cells, and targeted ES cell clones were identified by digestion of genomic DNA with BamHI, followed by a Southern blot analysis of ES cell DNA using a probe as indicated (Fig. 1B). Two Wdr34-targeted ES cell clones were injected into C57BL/6 blastocysts to generate chimeric founders, which were then bred with C57BL/6 to establish F1 heterozygotes. The Wdr34 heterozygotes were maintained in 129/SVE, C57BL/6, and SW mix background. PCR (polymerase chain reaction) analysis was used for routine genotyping with the following primers: forward primer BW1221F, 5’-TACCTCACTGCAGCTCTCTCA-3’ and reverse prime BW1221R, 5’-GTTGTACATCACCTGCAGGGA-3’ for the wild type allele, which produced a 200 bp fragment; and forward primer BW294, 5’-ATTGGGAAGACAATAGCAGGCA-3’and reverse primer BW1221R for the targeted Wdr34 allele, which produced a 280 bp fragment.

Cell lines and cell culture

Wild type and mutant Wdr34 primary mouse embryonic fibroblasts (pMEFs) were prepared from E11.5 or E12.5 mouse embryos. The procedure was approved by the Institutional Animal Care and Use Committee at Weill Medical College. The pMEFs were cultured in DMEM supplemented with 10% FBS (fetal bovine serum), penicillin, and streptomycin.

cDNA constructs, cloning, and transfection

To create GFP-Wdr34 or its mutant cDNA constructs, EGFP was first inserted into pLNCX retroviral vector (Clontech) by PCR and general cloning techniques. Mouse Wdr34 cDNA was amplified from a mouse cDNA library and cloned in-frame into pLNCX-EGFP. Wdr34-1-220 (amino acid residues) and Wdr34-221-E (221aa to the C-terminal end) fragments were amplified and cloned in-frame into the same vector. The constructs were verified by DNA sequencing. Virus carrying GFP, GFP-Wdr34, or its mutant constructs were generated by cotransfecting each of the viral constructs with pEco construct into Phoenix-Eco cells (ATCC) using the calcium phosphate precipitation method (31).

To generate MEFs that stably expressed GFP-Wdr34, GFP-Wdr34-1-220, or GFP-Wdr34-221-E, wild type MEFs were transduced with the above virus. About 36 hours after transduction, the cells were incubated in the medium with G418 (1 mg/ml) until neomycin resistant clones emerged. Heterogeneous stable cells were maintained in the medium with 0.1 mg/ml G418.

Embryo section immunofluorescence and whole mount in situ hybridization

For immunofluorescence of neural tube sections, mouse embryos at 10.5 days post coitus (E10.5) were dissected, fixed in 4% paraformaldehyde (PFA)/PBS for 1 h at 4 °C, equilibrated in 30% sucrose/PBS overnight at 4 °C, and embedded in OCT. The frozen embryos were transversely cryosectioned at the forelimb areas (10 μm/section). Tissue sections were immunostained using antibodies against Shh, Foxa2 (concentrated), Nkx2.2, Hb9, Isl1/2, Pax6, and Pax7 (Developmental Study Hybridoma Bank (DSHB), Iowa), or Arl13b and acetylated αtubulin together as described (41). Whole mount in situ hybridization of mouse embryos with digoxigenin-labeled riboprobes was performed as described (43).

Cell immunofluorescence and microscopy

For cell ciliation studies, cells were plated on coverslips coated with 0.1% gelatin for at least overnight and serum starved with 0.1% FBS for 24 h to arrest the cells. To assess Smo, Gli2, and Gli3 ciliary localization in response to Hh signaling, MEFs were incubated with control or ShhN conditioned medium (1:10 dilution with DMEM medium, ShhN working concentration ≥ 5 nM, 1% FBS final concentration) overnight before immunostaining. ShhN conditioned medium was collected from HEK293 cells transfected with pRK-ShhN expression vector. ShhN level was initially estimated by Western blot, and its activity was initially determined by Gli-luciferase reporter assay using Shh-light cells (44). For centrosome staining, cells were fixed in −20 °C cold methanol for 5 min. For the cytoplasmic and cilia staining, cells were fixed in 4% PFA/PBS for 15 min. After wash with PBS, the cells were incubated with blocking solution (PBS/0.2% Triton X-100/4% heat inactivated calf serum) for 20 min. The cells were then incubated with primary antibodies in blocking solution for 1 h at room temperature. The cells were washed with PBS, incubated with secondary antibodies in blocking solution for 1 h at room temperature. After wash three times with PBS, the coverslips were mounted to glass slides with Vectashield mounting fluid with DAPI (Vector Labs). The staining was visualized using a Zeiss Axiovert fluorescent microscope.

Antibodies

Arl13b antibodies were generated by Covance, Inc. Rabbits were immunized with an insoluble His-tagged mouse Arl13b purified from bacteria. It was used in a 1:1000 dilution. Other antibodies include: Smo (1:1000) (11), Gli2, Gli3, Ift88, Cep164, Odf2, Ta3, Cep290, GFP, Dnchc2, Lic3 (all 1:1000) (31,37,39), acetylated tubulin (1:4000), γ−tubulin (1:4000) (Sigma), Ift57 (1:1000), and IFT40 (1:1000) (45). Secondary antibodies Alexa Fluor 488-conjugated donkey anti-rabbit IgG and Cy3-conjugated donkey anti-mouse IgG were purchased from Jackson Immunoresearch, Inc.

Immunoblotting

E10.5 mouse embryos were washed with PBS and lysed in RIPA buffer (50 mM Tris-HCl [pH 7.4], 150 mM NaCl, 1 mM EDTA, 1%Triton X-100, 1% sodium deoxycholate, 0.1% SDS, protease inhibitors) for 10 min on ice. After being cleared by centrifugation, the lysates were mixed with protein loading buffer, resolved in SDS polyacrylamide gel, and transferred to a nitrocellulose membrane followed by western blotting as described (5).

Scanning electron microscopy

E9.5 wild type and Wdr34 mutant embryos were fixed with 2% paraformaldehyde/2% glutaraldehyde in 0.075 M sodium cacodylate buffer (EM quality) overnight at room temperature. After wash with PBS, the embryos were dissected to expose the lumen of the neural tube. The embryo fragments were dehydrated using a series of graded ethanol and subjected to critical point drying. The embryo fragments were mounted with the lumen faced-up on aluminum stubs with adhesive tabs and sputter-coated with gold-palladium alloy. Cilia of neural epithelia in the neural tube were imaged using a field emission electron microscope (Supra 25; Carl Zeiss).

Microtubule regrowth assay

MEFs were seeded onto coverslips coated with 0.1% gelatin in growth medium. After 24 h, they were treated with 1μg/ml nocodazole for 1h at 37 °C. The cells were washed once, incubated with normal growth medium to allow microtubule regrow and then fixed at 0, 5, 10, 20, and 60 min after nocodazole treatment, using −20 °C cold methanol. The cells were stained with αtubulin and DAPI and imaged.

Acknowledgements

We thank Dr. Gregory Pazour for Ift57 and Ift140 antibodies, Dr. Rajat Rohatgi for Smo antibody, Dr. Richard Vallee for Dnchc2 and Lic3 antibodies. We also thank Ms. Nina Lampen for assistance with SEM. Shh, Isl1, Hb9, Pax6, Pax7, Nkx2.2, and Foxa2 monoclonal antibodies were purchased from the Developmental Studies Hybridoma Bank maintained by the University of Iowa, Department of Biological Sciences, Iowa City, Iowa 52242, under contract NO1-HD-7-3263 from the NICHD. Procedures involved in mice were performed under an approved animal protocol. B.W. was supported by a National Institutes of Health, USA and K.T. and C.W. were supported by the Chinese Natural Science Foundation.

Conflict of Interest statement. None declared.

Funding

National Institutes of Health, USA (grant R01GM114429) and Chinese Natural Science Foundation (grant 81172294 and 81600401).

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