Many RNA-seq datasets have sought to examine genes that are enriched in cranial neural crest cells compared with other tissues (Simões-Costa et al., 2014) or axial levels (Martik et al., 2019). However, here we aimed to identify highly expressed transcription factors and chromatin remodelers that may have been missed due to overlapping expression between neural crest cells and surrounding tissues. To this end, gastrula stage Hamilton Hamburger (HH) four embryos were electroporated with the neural crest enhancer FoxD3-NC2:eGFP and cultured ex ovo until stage HH12 (Hamburger and Hamilton, 1951). The NC2 enhancer labels early migrating neural crest cells (Simões-Costa et al., 2012), thereby facilitating dissection of the region surrounding the rhombomere (r) six migratory neural crest stream for dissociation (Figure 1A–A’). To aid downstream analysis and clustering, we introduced an ‘outgroup’ of dissected primary heart tube cells into the single-cell suspension and generated barcoded Gel Bead-In-Emulsions (GEMs) on the 10X Genomics platform. The library was sequenced at a depth of 50,000 median reads/cell to profile a total of 1268 cells, out of which 1241 cells passed the quality control filters (Figure 1—figure supplement 1A–C).

Following mapping and dimensional reduction, the cells split into distinct cellular subtypes (Figure 1B), including five cell types (mesoderm, otic, ectoderm, hindbrain, and neural crest) derived from the dissected tissue, and the spiked-in outgroup (‘Heart tube’; Myl2⁺, Tnnt2⁺). Known genetic markers that were enriched in each population served to distinguish the neural crest subcluster (Tfap2B⁺, ItgB3⁺) from the surrounding tissues (i.e. otic placode (Cldn3⁺, Gbx2⁺); hindbrain (Pax6⁺, Zic2⁺); ectoderm (Epcam⁺, Crabp2⁺); mesoderm (FoxC2⁺, Col1A1⁺)) (Figure 1—figure supplement 2A–B). Consistent with the anatomical diversity of the mesoderm, the latter was further subdivided into specific cell types like myocardium (Hand2⁺) and paraxial mesoderm (Prrx1⁺) (Figure 1—figure supplement 1D–E). We particularly focused our subsequent analysis on the neural crest cluster in the context of the neighboring tissues of hindbrain and ectoderm (Figure 1C,D,D’).

We sought to determine all transcription factors and chromatin regulators that were expressed in the neural crest-specific subcluster, regardless of their expression in other cell types. To this end, we shortlisted all genes associated with the gene ontology terms ‘DNA-binding’, ‘regulation of transcription’, and ‘transcription factor’. This revealed several chromatin remodelers and transcription factors with high levels of expression in neural crest cells (Figure 1E; Figure 1—figure supplement 2C). The identified genes fell into two groups, the first of which was comprised of transcription factors enriched in neural crest cells, with little overlap in surrounding cell types. As expected, many of these genes, including Sox10, Ets1, MafB, and Nrip1, are known for their expression in the neural crest (Gandhi et al., 2020; Tani-Matsuhana et al., 2018). Importantly, the second group was comprised of chromatin remodelers and/or transcriptional regulators previously overlooked in bulk transcriptomic datasets, including Hmga1, Dact2, Ssrp1, and Tbxl1x, due to overlapping expression in other tissues. Indeed, their distribution in low-dimensional space confirmed that a high proportion of cells in the hindbrain and/or ectoderm also expressed these genes (Figure 1F). The expression of a subset of the above genes was validated at HH12 (Figure 1G–K) by in situ hybridization chain reaction (HCR), which revealed an abundance of transcripts in both r4 and r6 neural crest streams that emerge from the hindbrain. Taken together, the results show that our single-cell gene expression analysis is sufficient to resolve the underlying heterogeneity of the chick hindbrain. We also identified several genes expressed in migrating neural crest cells not highlighted in previous datasets given their broad expression in other tissues.

Of the novel transcriptional regulators that were previously overlooked in bulk transcriptomic datasets, we were particularly intrigued by the chromatin remodeler Hmga1, due to its extensively studied role in tumorigenesis. A member of the High Motility Group A (HMGA) superfamily, Hmga1 encodes a small, nonhistone chromatin remodeling protein that binds to the minor groove of DNA, thereby affecting the chromatin landscape and facilitating the binding of other transcription factors in the opposing major groove (Grosschedl et al., 1994). In developing mouse embryos, Hmga1 has been shown to have widespread expression across several tissues, including the brain, where its loss has been correlated with reduced developmental potential of neural precursor cells (Kishi et al., 2012). While well-studied in cancer (Masciullo et al., 2003; Sarhadi et al., 2006), little was known about its developmental function in neural crest cells. Given the parallels between the mechanisms that regulate delamination, migration, proliferation, and survival of neural crest cells and tumor cells (Gallik et al., 2017), we sought to characterize the expression of Hmga1 during neural crest development.

At HH12, Hmga1 was observed in migrating neural crest streams emanating from rhombomeres 4 and 6 (Figure 1K), and in the cranial mesenchyme, suggesting that its expression is not restricted to the hindbrain neural crest. Therefore, to determine its spatiotemporal pattern at early stages of neural crest development, we performed HCR for Hmga1 at stages ranging from gastrulation (HH4), when neural crest cells are undergoing induction in the neural plate border (Figure 2B’’’), to HH10, when neural crest cells have delaminated from the dorsal neural tube (Figure 2E’’’) and are mid-migration in the cranial region (Figure 2F’’’). As an early marker for the neural plate border and neural crest (Basch et al., 2006), we co-labeled Pax7 transcripts at the aforementioned stages.

Hmga1 transcripts were first detected in the neural plate and neural plate border, but not in the non-neural ectoderm at HH4+, and preceded the expression of Pax7 in the neural plate border. Hmga1 expression remained high at HH5-6 (Figure 2A–B), overlapping in the neural plate border (Figure 2—figure supplement 1A) with Pax7 transcripts (Figure 2B’), as observed in transverse sections (Figure 2B”, Figure 2—figure supplement 1B–B’). As the neural plate border elevated to form neural folds between HH7 and HH8 (Figure 2D’’’), expression of Hmga1 was retained in the neural tube (Figure 2C–D) and continued to overlap with Pax7 in the dorsal neural folds (Figure 2D’, Figure 2—figure supplement 1C–D’). Between stages HH9 and HH10, when neural crest cells delaminated from the dorsal neural tube (Figure 2E’’’) and started migrating laterally (Figure 2F’’’), Hmga1 expression was retained in delaminating (Figure 2E,E’,E’’, Figure 2—figure supplement 1E–F’) and migrating (Figure 2F,F’,F’’, Figure 2—figure supplement 1G–H’) neural crest cells. Interestingly, transverse sections through a representative HH10 embryo revealed that within the migrating neural crest stream, Hmga1 was expressed in both leader and follower cells, as compared to Pax7, which appeared to be downregulated in the leader cells (Figure 2F’’). Together, these results show that the onset of Hmga1 in the neural crest occurs in precursors at the neural plate border region prior to their specification and is retained in premigratory and migrating neural crest cells.

Given that Hmga1 transcripts were enriched in the cranial neural crest, we sought to interrogate its possible role therein. To this end, we designed guide RNA plasmids (gRNAs) targeting the coding sequence of Hmga1 (Figure 3—figure supplement 1A) and electroporated them together with constructs encoding Cas9 and nuclear RFP on the right side of HH4 gastrula stage embryos (Figure 3A). The left side of the embryo was electroporated with Cas9, nuclear RFP, and a control gRNA, chosen for its lack of binding in chick cells (Gandhi et al., 2017). Embryos were cultured ex ovo until stage HH9/9+ (Figure 3B), after which they were processed for immunohistochemistry, in situ hybridization, and HCR.

We first validated our knockout approach by probing for the expression of Hmga1 itself in knockout embryos using HCR. This revealed a significant reduction in the abundance of Hmga1 transcripts on the knockout side (Figure 3C). We quantified this phenotype in whole-mount embryos and observed a 25% reduction in Hmga1 expression (Figure 3D, Figure 3—source data 1; p<0.05, Wilcoxon rank test). Notably, the loss of Hmga1 transcripts in the neural crest was more dramatic than in the neural tube, due to targeted electroporation of knockout reagents to the presumptive neural plate border region. Next, we investigated the effect of knocking out Hmga1 on Pax7 expression in neural crest cells. We examined Pax7 mRNA expression by HCR in embryos where Hmga1 was knocked out on the right side, relative to the left side which served as an internal control. Consistent with their hierarchical onset of expression, loss of Hmga1 resulted in a notable reduction in Pax7 mRNA levels (Figure 3E). Next, we assessed whether the reduction in Pax7 transcripts would consequently result in a loss of Pax7 protein in the neural crest by immunostaining Hmga1-knockout embryos with a Pax7 antibody. As expected, Pax7 protein levels were dramatically reduced in the migratory cranial neural crest (Figure 3F,H’), with further analysis revealing a significant decrease in the number of Pax7+ cells on the knockout side compared to the control side (Figure 3G, Figure 3—source data 2; p<0.0001, student’s t-test). Moreover, in the absence of Hmga1, cranial neural crest cells failed to migrate properly, as depicted by the expression of the migratory neural crest marker HNK1 (Figure 3—figure supplement 1B). A transverse section through the hindbrain (Figure 3H) of another representative Hmga1-knockout embryo stained for Pax7 (Figure 3H’) revealed a notable reduction in the expression of Pax7 (Figure 3I) as well as the neural crest specifier Snail2 on the knockout side (Figure 3I’). Furthermore, using in situ hybridization, we found that other neural crest specifier genes including FoxD3 (Figure 3J,J’, Figure 3—figure supplement 1D,E’,E’’), Tfap2b (Figure 3K), Sox10 (Figure 3L,L’), and c-Myc (Figure 3—figure supplement 1C) were also downregulated on the knockout side. In transverse sections through Hmga1-knockout embryos labeled for Sox10 expression (Figure 3L), we also detected fewer Pax7+ cells (Figure 3N) and diminished levels of HNK1 expression (Figure 3M). On the other hand, no notable difference in the thickness of the neural tube was observed (Figure 3H,J’,M’’, Figure 3—figure supplement 1E), suggesting that the targeted knockout of Hmga1 in the neural plate border affected the neural folds/dorsal neural tube but not the neural plate itself. Taken together, our results indicate that Hmga1 is important for proper specification of neural crest cells.

As the expression of Hmga1 precedes that of Pax7 in the neural plate border, and its loss also causes a reduction of Pax7 levels in neural crest cells (Figure 3F), we next asked if this regulation occurs in the neural crest precursors that reside in the neural plate border. In the preceding experiments, we characterized the expression of neural crest markers following CRISPR-plasmid-mediated loss of Hmga1 at stages corresponding to early stages of neural crest emigration and migration. However, one caveat to our plasmid knockout strategy is that it takes time after electroporation of the CRISPR constructs in gastrula stage embryos for Cas9 to transcribe, translate, and properly fold to form a functional Cas9-gRNA complex. As specification of neural crest cells at the neural plate border is ongoing at HH4, this means that functional Cas9 would not be available until well after initial electroporation. Given that neural crest development occurs in a rostral-to-caudal wave along the body axis (Gandhi and Bronner, 2018), we speculated that effects of plasmid electroporation would be more penetrant in the hindbrain compared to anterior regions of the embryo, such as the midbrain. To test this possibility, we generated transverse sections through the midbrain (Figure 4A) and hindbrain (Figure 4B) of Hmga1-knockout embryos and quantified the number of Pax7+ cells as a ratio of cell count on the experimental versus control sides. Accordingly, we noticed that while this ratio was 0.948 ± 0.036 (n = 5) at the midbrain level, it was significantly reduced in the dorsal hindbrain, with a mean ratio of 0.69 ± 0.08 (n = 5; p<0.05, paired Student’s t-test) (Figure 4C, Figure 4—source data 1). These results supported our assumption that specification may have already occurred at the midbrain level by the time Cas9 was functionally active.

To circumvent this issue and test the earliest effects of knocking out Hmga1 concomitant with its onset of expression, we turned to an alternative CRISPR knockout strategy that enabled the loss of Hmga1 immediately after transfection. To this end, we electroporated recombinant Cas9 protein with in vitro-transcribed Hmga1 or control gRNA as ribonucleoprotein (RNP) complexes on the right and left sides of gastrula stage embryos, respectively (Figure 4D), and cultured embryos ex ovo until HH6. First, to validate the Cas9-protein-mediated knockout strategy, we labeled Hmga1 transcripts in knockout embryos using HCR and observed a very efficient reduction in Hmga1 expression (Figure 4E), especially in the neural plate border, thereby offering precise temporal control over the loss of this gene’s activity. Next, we assayed for the expression of Pax7 in the neural plate border by immunostaining Hmga1-knockout embryos and found that the levels of Pax7 protein in the neural plate border (Figure 4F, Figure 4—figure supplement 1A) were severely downregulated. Transverse sections through the experimental compared with control sides revealed that neural plate border cells no longer expressed Pax7 after the loss of Hmga1 (Figure 4H,I), and that this was not a result of premature apoptosis or aberrant cell proliferation (Figure 4—figure supplement 1B–G’, Figure 4—figure supplement 1—source data 1). In further support of the latter, the thickness of the neural plate border remained unchanged (Figure 4H’,I’). We also quantified the corrected Pax7 total fluorescence intensity (C.T.C.F) in the neural plate border and observed a statistically significant difference between the control and knockout sides (p<0.01, paired student’s t-test), with mean Pax7 intensity on the control side being 89.853 ± 23.388 a.u. (n = 5) as compared to 42.763 ± 16.079 a.u. (n = 5) on the knockout side (Figure 4J, Figure 4—figure supplement 1H, Figure 4—source data 2). Taken together, these results suggest that Hmga1 is required for the expression of Pax7 in neural crest precursors that are induced in the neural plate border.

The neural plate border was initially thought to contain discrete domains corresponding to neural, neural crest, placodal, and epidermal precursors. However, recent work has demonstrated that cells in this region co-express genes characteristic of different cell fates and exhibit a broad developmental potential, suggesting they are not restricted to individual cell fates until later in development (Roellig et al., 2017). In chick embryos, Tfap2a and Msx1 are expressed in the neural plate border, with Tfap2a transcripts spanning both the neural plate border and the non-neural ectoderm (de Crozé et al., 2011; Luo et al., 2003), whereas Msx1 transcripts are expressed within a subset of Pax7+ cells in the neural plate border region (Khudyakov and Bronner-Fraser, 2009). Given that loss of Hmga1 resulted in a reduction in Pax7 protein levels in the neural plate border, we asked if this was a result of a general neural plate border defect versus a selective effect on Pax7. To test this, we examined the expression of Tfap2a (Figure 4K) and Msx1 (Figure 4N) transcripts together with Pax7 using HCR in Hmga1-knockout embryos developed to neurula stages. Consistent with the loss of Pax7 protein in the neural plate border, Hmga1 knockout caused reduced Pax7 mRNA levels on the experimental (Figure 4M,P) compared to the control (Figure 4L,O) sides. However, the expression of Tfap2a (Figure 4L’,M’) and Msx1 (Figure 4O’,P’) was retained in the absence of Hmga1, together with no noticeable difference in the thickness of the neural plate border (Figure 4L’’,M’’,O’’,P’’). Since Pax7 and Msx1 are specifically expressed in the neural plate border, we quantified the corrected total cell fluorescence intensity per unit area associated with their transcripts and calculated the ratio between the experimental and control sides. Under control conditions, this ratio would be close to 1. However, for Pax7, the mean calculated ratio was 0.552 ± 0.06 (n = 6), with a statistically significant difference between the experimental and control sides (p<0.001, paired Student’s t-test). On the other hand, while the mean calculated ratio for Msx1 was 0.809 ± 0.098 (n = 3), the fluorescence intensities were not significantly different between the two sides (Figure 4Q, Figure 4—source data 3). Taken together, these results show that Hmga1 specifically regulates Pax7 expression at the neural plate border.

Given that loss of Hmga1 affects neural crest specification, we asked if its overexpression would have the converse effect. To exogenously provide Hmga1, we designed a plasmid construct containing the coding sequence of Hmga1 under the regulation of a ubiquitous enhancer/promoter combination (Figure 5A). This construct also contained the coding sequence for nuclear RFP downstream of an Internal Ribosome Entry Site (IRES), allowing identification of successfully transfected cells. To test the effect of overexpressing Hmga1, we electroporated this construct on the right side of gastrula stage embryos and cultured them ex ovo until HH9+. The left side served as an internal control and was electroporated with an equal concentration of a construct encoding nuclear RFP alone (Figure 5B). The results show that, rather than having the opposite effect to loss of function, overexpression of Hmga1 also resulted in a notable reduction in Pax7 expression on the experimental side (Figure 5C). This suggests that maintaining appropriate levels of Hmga1 is critical for proper neural crest specification.

The gold standard to demonstrate specificity for loss-of-function experiments is to perform a rescue. We posited that if modulating the levels of Hmga1 was important for neural crest formation, then exogenous expression of Hmga1 in an Hmga1-knockout background should successfully rescue neural crest cell numbers. Since the Protospacer Adjacent Motifs (PAMs) adjacent to both Hmga1 gRNAs are located in the introns (Figure 3—figure supplement 1A), the coding sequence on the plasmid would be guarded against the endonuclease activity of Cas9. To test our hypothesis, we knocked out Hmga1 using CRISPR plasmids as described above, but co-electroporated the ‘rescue’ construct on the right side. The left side was electroporated with an equal concentration of a plasmid encoding nuclear RFP. Embryos cultured to HH9+ and immunostained for Pax7 revealed that overexpression of Hmga1 concomitant with knocking out the endogenous gene successfully rescued Pax7 levels in cranial neural crest cells (Figure 5D). To quantify the extent of rescue, we calculated the ratio of the number of Pax7+ cells on the experimental versus control sides in both wholemount ‘knockout’ (from Figure 3G) and ‘rescue’ embryos (Figure 5D). In unperturbed embryos, this ratio will be close to 1, reflecting similar numbers of Pax7+ cells on both sides of the embryo. However, in the ‘knockout’ group, we observed a mean ratio of 0.767 ± 0.041 (n = 11), which was significantly different from the ratio of 0.946 ± 0.016 (n = 8) observed in the ‘rescue’ group (p<0.001, Welch two-sample t-test) (Figure 5E, Figure 5—source data 1). Next, we probed for the expression of the neural crest specifier gene Sox10 to ask if rescuing the expression of Hmga1 truly restored the process of neural crest specification. To do this, we processed ‘rescue’ embryos for HCR against Sox10 (Figure 5F) together with Pax7 (Figure 5G). Indeed, restoring the levels of Hmga1 was sufficient to rescue the expression of Sox10 (Figure 5G’) and Pax7 (Figure 5G’’) in early migrating crest, and Pax7 in the premigratory crest residing in the dorsal neural tube, as visualized in transverse sections through the embryo. We also confirmed that the expression of the ‘rescue’ construct was restricted to the dorsal neural tube (Figure 5G’’’), thereby precluding unintended effects on neural tube development.

Finally, given that the loss of Hmga1 specifically affected Pax7 expression in neural crest precursors, we asked if exogenous expression of Pax7 would be sufficient to rescue the effects of losing Hmga1 on neural crest specification. We tested this by overexpressing the coding sequence of Pax7 (Roellig et al., 2017) on the right side of gastrula stage embryos together with CRISPR plasmids targeting Hmga1 (Figure 5H,H’). Given that the effect of CRISPR-plasmid-mediated loss of Hmga1 was more penetrant posteriorly, we developed ‘Pax7-rescue’ embryos to HH9+, processed them for HCR against the neural crest specifier genes Tfap2b (Figure 5I) and Snai2 (Figure 5I’), and generated transverse sections through the hindbrain. Tfap2b is expressed in delaminating and migrating neural crest cells (Simoes-Costa and Bronner, 2016), whereas Snai2 is expressed in premigratory neural crest and is eventually downregulated as the cells begin to migrate (Taneyhill et al., 2007). Compared to the Hmga1-knockout embryos in which Tfap2b mRNA (Figure 3K) and Snail2 protein (Figure 3I’) levels were notably downregulated, restoring the levels of Pax7 in an Hmga1 knockout background rescued the expression of both Tfap2b (Figure 5J) and Snai2 (Figure 5K) in the dorsal hindbrain. Together, these results suggest that maintaining the correct levels of Hmga1 is necessary for proper neural crest specification in a Pax7-dependent manner.

Neural crest induction, specification, and emigration from the neural tube are intricate processes that require an interplay between Wnt, FGF, and BMP signaling pathways (Piacentino and Bronner, 2018; Woda et al., 2003) working reiteratively at different stages of development. For example, at the onset of neural crest emigration, Wnt1 is prominently expressed in the dorsal neural tube, where premigratory neural crest cells reside (Simões-Costa et al., 2015). As a result, these cells turn on Snai2, a critical regulator of EMT (Nieto et al., 1994) known to function downstream of the Wnt signaling pathway.

After knocking out Hmga1 using CRISPR plasmids, we noted not only perturbed emigration but also a dramatic reduction in Snail2 levels, even within the subset of Pax7-expressing cells in the dorsal neural folds (Figure 3I’). These results raised the intriguing possibility that this might be due to an effect on Wnt signaling in already-specified premigratory neural crest cells. Accordingly, we hypothesized that Hmga1 may function as a Wnt activator in these cells. If so, its loss would be predicted to result in reduced Wnt signaling in the dorsal neural tube. To test this possibility, we used a reporter construct expressing nuclear GFP under the regulation of six Tcf/Lef binding sites and a minimal promoter as a readout for canonical Wnt signaling (Ferrer-Vaquer et al., 2010; Figure 6A). Plasmids encoding Cas9, gRNAs targeting Hmga1, nuclear RFP, and Tcf/Lef:H2B-GFP were electroporated on the right side of gastrula stage embryos, while the left control side was electroporated with plasmids encoding Cas9, control gRNA, nuclear RFP, and Tcf/Lef:H2B-GFP. As described above, this plasmid-based knockout strategy resulted in the loss of Hmga1 after neural crest specification in the midbrain but well before their emigration. Embryos were allowed to develop until HH9, by which time neural crest cells have started delaminating from the neural tube at the midbrain level (Figure 6B). Consistent with our hypothesis, the results show that Hmga1 knockout caused a notable reduction in canonical Wnt reporter activity on the knockout side compared to the control side (Figure 6C) at the midbrain level. Interestingly, these embryos had a neural crest migration defect (Figure 6—figure supplement 1C–D) but no notable difference in the number of Pax7+ cells between the two sides (Figure 6—figure supplement 1E’), as expected if the Hmga1 knockout occurred after specification was complete; this is consistent with previous work showing that perturbation of canonical Wnt signaling following specification does not affect the number of Pax7+ cells at cranial EMT stages (Hutchins and Bronner, 2018). Quantitation of this effect revealed a significant reduction in reporter activity following the loss of Hmga1 (Figure 6D, Figure 6—source data 1; p<0.01; student’s t-test), as measured by comparing the ratio between the knockout and the control sides of transfected cells (RFP+) that turned on Wnt signaling within the Pax7+ domain, therefore expressing GFP. While this ratio was expected to be one for embryos with unperturbed Wnt signaling on both sides, we observed a mean ratio of 0.325 ± 0.082 (n = 5), suggesting that Wnt activity was disrupted in the absence of Hmga1.

Next, to investigate the mechanism by which Hmga1 regulates Wnt signaling, we turned to a recently published cranial neural crest chromatin accessibility dataset (Williams et al., 2019) and looked for open chromatin regions surrounding genes that encode for known Wnt ligands. In particular, Wnt1 expression in the dorsal neural tube is known to be necessary for proper delamination of cranial neural crest cells (Simões-Costa et al., 2015). Interestingly, we discovered a putative enhancer downstream of Wnt1 (Figure 6—figure supplement 1A) that contained an AT-rich domain consistent with Hmga1-binding motifs (Figure 6—figure supplement 1B; Reeves and Nissen, 1990). Therefore, we hypothesized that Hmga1 may modulate Wnt signaling by regulating Wnt1 expression. To test this, we knocked out Hmga1 on the right side of gastrula stage embryos using CRISPR plasmids, cultured them ex ovo until HH9, and examined Wnt1 mRNA expression using in situ hybridization. Indeed, the dorsal neural tube expression of Wnt1 was severely downregulated (Figure 6E) in the midbrain. Consistent with the effect of losing Hmga1 after neural crest specification, the number of Pax7+ cells in the dorsal neural tube appeared unchanged (Figure 6E’). In contrast, no change in Wnt1 expression was observed at the hindbrain level (Figure 6F) which, being developmentally ‘younger,’ instead exhibited a specification defect that resulted in fewer Pax7+ cells in the dorsal neural tube on the experimental side compared to the control side (Figure 6F’). Interestingly, following Hmga1 knockout, we also observed defects in basement membrane remodeling and laminin channel formation at midbrain levels (Figure 6G), another Wnt-dependent process necessary for neural crest EMT; consistent with Wnt inhibition via Draxin overexpression (Hutchins and Bronner, 2019), loss of Hmga1 abrogated laminin remodeling and resulted in physical blockage of the channel through which migrating cranial neural crest cells normally transit (Figure 6H). Together, these data indicate that after neural crest specification, Hmga1 is necessary for the expression of Wnt1 and activation of canonical Wnt signaling in the dorsal neural tube, and by extension, Wnt-dependent neural crest delamination/EMT.

Finally, given that Hmga1 functions as a canonical Wnt pathway activator, we asked if the migration defects caused by the loss of Hmga1 post-specification can be rescued by restoring canonical Wnt signaling in premigratory neural crest cells. To address this, we expressed GFP-tagged, stabilized ß-catenin (NC1-∆90 ß-cat) to upregulate canonical Wnt signaling output specifically in premigratory neural crest cells, thus circumventing the critical process of neural crest induction at the neural plate border (Hutchins and Bronner, 2018). If loss of Hmga1 in premigratory neural crest cells resulted in migration defects due to reduced canonical Wnt signaling, expression of a stabilized ß-catenin would be predicted to restore those levels, thereby rescuing proper migration. To test this, we knocked out Hmga1 on the right side of gastrula stage embryos using CRISPR plasmids as previously described, but co-electroporated NC1-∆90ß-cat-GFP on the right side. The left side was electroporated with control reagents (Figure 6I). Embryos were cultured ex ovo until HH9+ and processed for immunohistochemistry against Pax7. Consistent with our hypothesis that Hmga1 functions as a Wnt activator in neural crest cells, expression of stabilized ß-catenin was sufficient to rescue proper cranial neural crest migration from Hmga1 knockdown (Figure 6J). We also calculated the ratio of the area occupied by migrating cranial neural crest cells between the experimental and control sides (Figure 6K, Figure 6—source data 2). For wildtype embryos, this ratio would be close to 1, reflecting equal neural crest migration on both sides of the embryo. However, after the loss of Hmga1, the ratio of areas on experimental versus control sides was 0.642 ± 0.028 (n = 5). Importantly, co-expression of stabilized ß-catenin in premigratory neural crest rescued migration, with an average migration ratio of 1.015 ± 0.035 (n = 4), which was significantly different from the knockout group (p<0.0001, ANOVA and post hoc Tukey HSD). Similarly, ubiquitous expression of the Hmga1 coding sequence also rescued neural crest migration, with an average migration ratio of 0.972 ± 0.044 (n = 5), which was also significantly different from the knockout group (p<0.001, ANOVA and post hoc Tukey HSD). Taken together, our results suggest that Hmga1 mediates the process of EMT by activating canonical Wnt signaling in premigratory neural crest cells, thus enabling them to emigrate from the neural tube.

Chicken embryos (Gallus gallus) were commercially obtained from Sun Valley farms (CA), and developed to the specified Hamburger-Hamilton (HH) (Hamburger and Hamilton, 1951) stage in a humidified 37°C incubator. For ex ovo electroporations, embryos were dissected from eggs at HH4, injected with specified reagents, then electroporated as described previously (Sauka-Spengler and Barembaum, 2008). Following electroporation, embryos were cultured in fresh albumin/1% penicillin-streptomycin at 37°C and grown to specified HH stages. Once the embryos reached the desired stages, they were screened for transfection efficiency and overall health. Unhealthy and/or poorly transfected embryos were discarded and not included for downstream assays.

HH4 embryos electroporated with FoxD3-NC2:eGFP were cultured until HH12 ex ovo at 37°C, following which the hindbrain region spanning rhombomeres 6, 7, and 8 was dissected under a fluorescence microscope. For dissociation, several different conditions were tested (dissociation in a glass dish for 1 hr in Accumax (EMD Millipore), chemical dissociation on a nutator for 15, 30, and 45 min, and chemical dissociation with gentle pipetting for 15, 30, and 45 min). The quality of the single-cell suspension obtained was tested by a trypan blue-based live-dead staining. Accordingly, we pooled dissected tissue washed in chilled 1X DPBS and incubated it in Accumax cell dissociation solution for 15 min at 37°C with gentle mixing every 5 min. Dissociation was terminated using Hanks Buffered Saline Solution (HBSS) (Corning) supplemented with BSA Fraction V (Sigma; 0.2% w/v). The suspension was centrifuged at 300 g for 4 min to collect cells at the bottom, the supernatant was removed, and the pellet was resuspended in 1 mL HBSS-BSA. To remove cell debris and clumps, the 1 mL suspension was passed through a 20 µm filter in a clean hood. This suspension was loaded on a 10X Chromium chip A (v2) to generate GEMs. The library was prepared according to the manufacturer’s protocol and sequenced on the Illumina HiSeq platform using the paired end chemistry.

The raw fastq files were aligned to the galgal6 (GRCg6a) genome assembly obtained from the ENSEMBL database using the cellranger pipeline downloaded from the 10X Genomics website. For feature counts, a custom galgal6 GTF file, where all annotated 3’ UTRs were extended by 1 kb, was used. This was done to compensate for improper gene annotations in the chick genome. The count matrices were then imported in R for analysis using Seurat (Butler et al., 2018). The initial filtering step discarded all cells with fewer than 200 and more than 10,000 genes per cell. We also filtered out cells expressing more than 5% mitochondrial or less than 20% ribosomal genes. Next, we removed genes corresponding to small RNAs, micro RNAs, mitochondria, and general housekeeping from the count matrix. Following log normalization and Principal Component Analysis, the cells were clustered using the first 15 dimensions (calculated from the elbow plot). Different resolution parameter values were tested, and a value of 0.45 was used to identify subpopulations within the data. Dimensional reductionality was performed using the UMAP (McInnes et al., 2018) algorithm. All plots were created in R, exported in SVG format, and assembled in Inkscape.

HCR v3 was performed using the protocol suggested by Molecular Technologies (Choi et al., 2018) with minor modifications. Briefly, the embryos were fixed in 4% paraformaldehyde (PFA) overnight at 4°C or 2 hr at room temperature, washed in 0.1% PBS-Tween, dehydrated in a series of 25%, 50%, 75%, and 100% methanol washes, and incubated overnight at −20°C in 100% methanol. The next day, the embryos were rehydrated, treated with proteinase-K for 2–2.5 min, and incubated with 10 pmol of probes dissolved in hybridization buffer overnight at 37°C. The next day, following several washes in ‘probe wash buffer,’ embryos were incubated in 30 pmol of hairpins H1 and H2 diluted in Amplification buffer at room temperature overnight. The next morning, embryos were washed in 0.1% 5x-SSC-Tween and imaged. All probes were designed and ordered through Molecular Technologies.

The coding sequence of Hmga1 was obtained from the UCSC genome browser (Karolchik et al., 2003). A V5 tag was cloned in-frame at the N-terminus using overlap PCR (Accuprime). The fusion product was cloned downstream of the CAGG promoter, upstream of the IRES-H2B-RFP segment of pCI-H2B-RFP (Betancur et al., 2010) to clone the final plasmid (CAGG >V5-HMGA1-IRES-H2B-RFP). The Cas9 and gRNA constructs (Gandhi et al., 2017), neural crest enhancer FoxD3-NC2:eGFP (Simões-Costa et al., 2012), canonical Wnt reporter TCF/Lef:H2B-GFP (Ferrer-Vaquer et al., 2010), and neural crest-specific stabilized ß-catenin NC1-∆90ß-cat (Hutchins and Bronner, 2018) have all been previously described and validated.

The genomic locus for Hmga1 was obtained from the UCSC genome browser (Karolchik et al., 2003). Two gRNAs targeting the coding sequence, the first targeting exon 3 (5’-CCAGGAAGAAACCGGAGgta-3’), and the second targeting exon 4 (5’-GCCAGCTCCAAAGGCAGGgt-3’), were designed using CHOPCHOP (Labun et al., 2019). The protospacers were cloned downstream of the chick U6.3 promoter as described in Gandhi et al., 2017. For control electroporations, the control gRNA described in Gandhi et al., 2017 was used. CAGG >nls-Cas9-nls and CAGG >H2 B-RFP were electroporated at a concentration of 2 µg/µl, together with either 0.75 µg/µl per Hmga1 gRNA on the right side or 1.5 µg/µl of control gRNA on the left side.

For Cas9/in vitro-transcribed gRNA RNP experiments, we generated single-guide RNAs (sgRNAs) as described previously (Hutchins and Bronner, 2018), using the following primers:

Of the recombinant Cas9 (M0646; New England Biolabs), 2.6 µl was mixed with equal volumes of control gRNA or 1.3 µl each of the two Hmga1 gRNAs, and heated to 37°C for 15 min. The solution was then incubated at room temperature for 15 min, mixed with 2 µg/µl H2B-RFP and 1 µl of sterilized 2% food dye, and injected in embryos for electroporation.

Chromogenic in situ hybridization was performed as described previously for FoxD3, Sox10, and Tfap2b, c-Myc, and Wnt1 (Kerosuo and Bronner, 2016; Simoes-Costa and Bronner, 2016; Simões-Costa et al., 2015).

Immunohistochemistry was performed as described previously (Gandhi et al., 2017). Briefly, embryos were fixed for 20 min at room temperature, blocked in 10% goat or donkey serum in 0.5% PBS-Triton overnight at 4°C, incubated overnight at 4°C in primary antibodies diluted in blocking solution, washed at room temperature in 0.5% PBS-Triton, incubated overnight at 4°C in secondary antibodies diluted in blocking solution, washed at room temperature in 0.5% PBS-Triton, and processed for imaging and/or cryosectioning. The following primary antibodies and concentrations were used: Mouse IgM HNK1 (1:5; Developmental Studies Hybridoma Bank (3H5)); Mouse IgG1 Pax7 (1:10; Developmental Studies Hybridoma Bank (RRID:AB_528428)); Goat GFP (1:500; Rockland Cat# 600-101-215); Rabbit RFP (1:500; MBL Cat# PM005); Rabbit Snail2 (1:200; Cell Signaling Technology (9585)); Rabbit Laminin (1:1000; Sigma-Aldrich (L9393)); Rabbit cleaved-Caspase 3 (1:500; R and D Systems Cat# AF835); Mouse phospho-histone H3 (1:500; Abcam Cat# Ab14955). The following species-specific secondary antibodies labeled with Alexa Fluor dyes (Invitrogen) were used: Goat/Donkey anti-Mouse Alexa Fluor 647 (for Pax7 and pH3; 1:250), Goat/Donkey Goat anti-Mouse IgM Alexa Fluor 350/488 (for HNK1; 1:250), Goat/Donkey anti-Rabbit Alexa Fluor 488 (for Snail2, cleaved-Caspase3, and Laminin; 1:250), Donkey anti-goat Alexa Fluor 488 (for Citrine; 1:500), and Goat/Donkey anti-rabbit Alexa Fluor 568 (for RFP; 1:500).

Following whole mount imaging, embryos were washed in 5% and 15% sucrose overnight at 4°C. The next day, embryos were transferred to molten gelatin for 3–5 hr at 37°C, embedded in molds at room temperature, frozen in liquid nitrogen, and stored at −80°C overnight. Embedded embryos were sectioned on a micron cryostat to obtain 16 µm sections through immunostained embryos and 20 µm sections through in situ hybridized embryos. The sections were degelatinized at 42°C in 1x PBS for 5 min, washed in 1x PBS, soaked in 1x PBS containing 0.1 µg/mL DAPI for 2 min, washed in 1x PBS and distilled water. Fluoromount mounting medium was used to mount coverslips on slides.

Whole mount embryos and sections on slides were imaged on a Zeiss Imager M2 with an ApoTome module and/or Zeiss LSM 880 confocal microscope at the Caltech Biological Imaging Facility. Images were post-processed using FIJI imaging software (Schindelin et al., 2012). To calculate corrected total cell fluorescence (CTCF), the following formula was used:CTCF=Integrated Density-(Selected area*Mean background fluorescence)

For cell counts, a median filter was applied to 8-bit images. A Bernsen-based auto local-thresholding method (Bernsen, 1986) followed by watershed segmentation was used to identify cell boundaries. The ‘Analyze particles’ function was used to count the number of cells. All statistical analyses were performed in R. The Wilcoxon rank test was used in datasets that were not normally distributed. In cases where the underlying distribution was normal, a student’s t-test was used to calculate significance. In cases where multiple samples were compared, Analysis of Variance (ANOVA) test combined with Tukey HSD correction was used. Post hoc power analysis was used to validate sample size and confirm sufficient statistical power (>0.8).