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Distinct expression patterns of Hedgehog signaling components in mouse gustatory system during postnatal tongue development and adult homeostasis

  • Archana Kumari ,

    Roles Conceptualization, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration, Resources, Supervision, Writing – original draft, Writing – review & editing

    Affiliation Department of Biologic and Materials Sciences & Prosthodontics, School of Dentistry, University of Michigan, Ann Arbor, Michigan, United States of America

  • Nicole E. Franks,

    Roles Investigation, Writing – review & editing

    Affiliation Department of Cell and Developmental Biology, Medical School, University of Michigan, Ann Arbor, Michigan, United States of America

  • Libo Li,

    Roles Investigation, Writing – review & editing

    Current address: Department of Pharmaceutical Sciences, College of Pharmacy, University of Michigan, Ann Arbor, Michigan, United States of America

    Affiliation Department of Biologic and Materials Sciences & Prosthodontics, School of Dentistry, University of Michigan, Ann Arbor, Michigan, United States of America

  • Gabrielle Audu,

    Roles Investigation, Writing – review & editing

    Affiliation Department of Cell Biology and Neuroscience, Rowan-Virtua School of Translational Biomedical Engineering and Sciences, Virtua Health College of Medicine and Life Sciences of Rowan University, Stratford, New Jersey, United States of America

  • Sarah Liskowicz,

    Roles Investigation, Writing – review & editing

    Affiliation Department of Biology, University of Scranton, Scranton, Pennsylvania, United States of America

  • John D. Johnson,

    Roles Investigation, Writing – review & editing

    Affiliation Rowan-Virtua School of Osteopathic Medicine, Virtua Health College of Medicine and Life Sciences of Rowan University, Stratford, New Jersey, United States of America

  • Charlotte M. Mistretta,

    Roles Conceptualization, Formal analysis, Funding acquisition, Methodology, Project administration, Resources, Supervision, Writing – original draft, Writing – review & editing

    Affiliation Department of Biologic and Materials Sciences & Prosthodontics, School of Dentistry, University of Michigan, Ann Arbor, Michigan, United States of America

  • Benjamin L. Allen

    Roles Conceptualization, Formal analysis, Funding acquisition, Methodology, Project administration, Resources, Supervision, Writing – review & editing

    Affiliation Department of Cell and Developmental Biology, Medical School, University of Michigan, Ann Arbor, Michigan, United States of America


The Hedgehog (HH) pathway regulates embryonic development of anterior tongue taste fungiform papilla (FP) and the posterior circumvallate (CVP) and foliate (FOP) taste papillae. HH signaling also mediates taste organ maintenance and regeneration in adults. However, there are knowledge gaps in HH pathway component expression during postnatal taste organ differentiation and maturation. Importantly, the HH transcriptional effectors GLI1, GLI2 and GLI3 have not been investigated in early postnatal stages; the HH receptors PTCH1, GAS1, CDON and HHIP, required to either drive HH pathway activation or antagonism, also remain unexplored. Using lacZ reporter mouse models, we mapped expression of the HH ligand SHH, HH receptors, and GLI transcription factors in FP, CVP and FOP in early and late postnatal and adult stages. In adults we also studied the soft palate, and the geniculate and trigeminal ganglia, which extend afferent fibers to the anterior tongue. Shh and Gas1 are the only components that were consistently expressed within taste buds of all three papillae and the soft palate. In the first postnatal week, we observed broad expression of HH signaling components in FP and adjacent, non-taste filiform (FILIF) papillae in epithelium or stroma and tongue muscles. Notably, we observed elimination of Gli1 in FILIF and Gas1 in muscles, and downregulation of Ptch1 in lingual epithelium and of Cdon, Gas1 and Hhip in stroma from late postnatal stages. Further, HH receptor expression patterns in CVP and FOP epithelium differed from anterior FP. Among all the components, only known positive regulators of HH signaling, SHH, Ptch1, Gli1 and Gli2, were expressed in the ganglia. Our studies emphasize differential regulation of HH signaling in distinct postnatal developmental periods and in anterior versus posterior taste organs, and lay the foundation for functional studies to understand the roles of numerous HH signaling components in postnatal tongue development.


The tongue houses taste buds in three different taste papillae at three different locations: fungiform papilla (FP) on the anterior two thirds, circumvallate papilla (CVP) on the posterior third mid-dorsum and foliate papilla (FOP) bilaterally on the posterior sides. These papilla types and resident taste buds develop with different time courses in mammalian species [13]. In sheep for example, the taste buds make a fetal appearance, continue development in utero and have extended postnatal maturation [4]. In rat and mouse, the lingual taste papillae emerge in the embryo; although initial taste buds-like cell clusters are seen before birth, taste buds per se do not emerge until postnatal stages [57]. However, both papillae and taste buds continue to develop during early postnatal stages and reach maturity in the anterior and posterior regions by the third and sixth postnatal weeks [1,810]. A mature FP harbors a single taste bud, while a CVP and multiple rows of FOP house numerous taste buds tightly associated with each other (Fig 1A). Anterior tongue and FP receive innervation from chorda tympani and lingual nerve fibers and posterior tongue and CVP are innervated by the glossopharyngeal nerve (Fig 1A). Interestingly, FOP anterior ridges receive chorda tympani nerve fibers and posterior ridges are also innervated by the glossopharyngeal nerve (Fig 1A) [1113]. The distinct location, morphology, and innervation of three lingual taste papillae may lead to their different trajectories of embryonic and postnatal development.

Fig 1. Schematic of lingual taste papillae and Hedgehog (HH) signaling.

(A) Illustrations of tissue architecture for the fungiform (FP), circumvallate (CVP) and foliate (FOP) papillae. FP is presented in a sagittal section, adjacent to spinous, non-taste filiform papilla. The CVP and FOP are illustrated in a horizontal section (to the dorsal tongue surface), and boxes refer to magnified taste bud (TB) distribution along the epithelium. Each TB is composed of taste cells and TB basal cells, and is surrounded by perigemmal cells. All the papillae comprise basal cells in the epithelium, extragemmal cells on the apical surface and stromal cells in the connective tissue core. Gustatory (chorda tympani and glossopharyngeal) and mechanosensory (lingual) nerves are indicated. FOP anterior (a) and posterior (p) ridges are indicated. (B) HH ligand binds PTCH1, releasing SMO inhibition to modulate GLI proteins and activate HH target gene expression. (C) HH signaling is regulated by the co-receptors, GAS1, CDON and BOC (green, activator), and the secreted antagonist HHIP (red, repressor). Transcriptional readouts of HH signaling are controlled by three GLI proteins: GLI1 (green, activator), GLI2 (light green, activator>repressor), and GLI3 (red, repressor>activator).

In our current study, we use mouse models. The number of mouse FP taste cells increases rapidly by postnatal day 7 (P7), more slowly between P7 and P14, and finally reaches a steady level by P21 [10,14,15]. In contrast, the number of rodent CVP and FOP taste buds increases threefold between P7 and P45 and reaches a maximum at P90 [1,10,16]. In addition, formation of taste pores, remodeling of gustatory innervation and enlargement of tongue muscles also occurs during the initial postnatal weeks [10,1618]. These various and extreme changes must be tightly coordinated to create a stereotypical, functionally mature taste organ.

Hedgehog (HH) signaling is an essential regulator of embryonic patterning [1921], postnatal organ development [22,23] and homeostasis [24] of several organs. We have previously shown the role of epithelial HH/GLI signaling in the maintenance of adult taste organs [25], and that HH signaling through Smoothened (SMO) also regulates papilla/taste buds homeostasis and neural responses [2628]. HH signaling is also active and vital during tongue development [5,7,29,30]. However, few studies have investigated HH signaling regulation in the anterior tongue during the initial postnatal weeks when the dynamic papillae and taste buds are continuously growing [8,10,15]. Further, HH signaling regulation in posterior CVP and FOP remain to be investigated. Thus, identification of signaling component activities during postnatal lingual development and maturation is essential to understand HH regulatory roles.

Fundamentally, HH signaling initiates through ligand binding to the canonical receptor Patched 1 (PTCH1), which relieves inhibition of SMO, leading to downstream signaling through the modulation of GLI proteins and altered transcription of HH pathway targets (Fig 1B) [3133]. Notably, HH pathway activation also requires ligand interactions with the co-receptors CDON, BOC and GAS1 [34], whereas the pathway is inhibited by binding of the HH antagonist HHIP to HH ligands (Fig 1C) [35,36]. Furthermore, HH transcription requires the combined use of three different GLI proteins. GLI1 is a transcriptional target and encodes an exclusive activator of the HH pathway; GLI2 is the major transcriptional activator; GLI3 acts principally as a repressor in HH signal transduction (Fig 1C) [37]. The field of taste research has mainly focused on the ligand, sonic HH (SHH), and the target gene, Gli1, while PTCH1, GLI2 and GLI3 have received less attention. Further, the obligatory membrane receptors GAS1, BOC and CDON and the HH antagonist HHIP remain largely unexplored. Our recent studies with HHIP [27] emphasize that these less studied HH pathway components might not be ‘secondary’ signaling elements but rather key signaling molecules that regulate organ integrity and thus need to be addressed.

As FP, CVP and FOP are distinct in their development and structural organization, their signaling regulation might be different. Thus, we investigated the expression pattern of HH signaling components in anterior FP and posterior CVP and FOP tissue regions: taste buds, basal, perigemmal, apical extragemmal and stromal cells (Fig 1A). We studied the ligand Shh, membrane receptors Ptch1, Gas1, Cdon and HH antagonist Hhip, and transcription factors Gli1, Gli2 and Gli3 in developing FP, CVP and FOP at the early postnatal stages between P3 and P10 (termed P7), and compared the pattern with that of mature taste papillae in mice aged eight weeks or older (termed adult). We also analyzed anterior FP during a later development stage between P21 and P31 (termed P28). In addition, we mapped HH pathway components in the adult soft palate, and geniculate and trigeminal ganglia with soma for chorda tympani and lingual nerve fibers, respectively, in the anterior tongue. We have used X-gal staining of lacZ- reporter mouse models or immunostaining to map the pathway components. The data reveal unique expression of the positive regulators of HH signaling activity, Ptch1, Gas1, Cdon, Gli1 and Gli2 in FP, CVP and FOP, and soft palate. Further differences are noted in the expression pattern of several HH signaling components in the anterior tongue, which is broader in the early postnatal stages as compared to adulthood. Notably, SHH, the receptor Ptch1, and the transcription factors Gli1 and Gli2 are the only components of the HH pathway observed in both geniculate and trigeminal ganglia. Overall, our data suggest a shift in HH pathway regulation of taste organs after the conclusion of postnatal tongue maturation.

Materials and methods


All animal use and care procedures were performed according to the guidelines of the National Institutes of Health. Informed written consent for all the protocols of the University of Michigan and Rowan University was obtained by the Institutional Animal Care and Use Committee (IACUC). Male or female mice, from the first postnatal week through adult stages were used. Pups less than 10 days of age were euthanized by decapitation, while in pups more than 10 days old, we used CO2 overdose, followed by decapitation.

Mouse models/strains

lacZ reporters.

Mice carrying lacZ alleles for the ligand (ShhlacZ/+, MGI:2678342) [38,39], HH-receptor Ptch1 (Ptch1lacZ/+, Jackson Laboratories strain:003081) [40], HH co-receptor Cdon (CdonlacZ/+) [41], HH antagonist Hhip (HhiplacZ/+, Jackson Laboratories strain:006241) [42], HH target gene/responding Gli1 (Gli1lacZ/+) (Jackson Laboratories strain:008211), HH transcriptional activator Gli2 (Gli2lacZ/+, Jackson Laboratories strain:007922) [43] and transcriptional repressor Gli3 (Gli3lacZ/+) [44] were maintained on a mixed 129S4/SvJaeJ/C57BL6/J background. HH co-receptor Gas1 (Gas1lacZ/+) [20] was maintained on a C57BL6/J background.

RFP reporter.

Mice carrying tamoxifen-inducible expression of RFP in Shh-expressing cells and their progeny (ShhCreERT2;R26RFP) were used. ShhCreERT2 (Jackson Laboratories strain:005623); R26RFP (Jackson Laboratories strain:007914) double transgenic animals were given 400 mg/kg of tamoxifen in Teklad Global Diet (Harlan) daily for 14–30 days.

Tissue dissection and processing

Tongues on mandibles were collected between postnatal day (P) 3 and P10 (termed P7), between P21 and P31 (termed P28) and between 8–15 weeks of age (termed adult). Soft palate (SP) and the ganglia (geniculate, GG and trigeminal, TG) were dissected at adult stages. All the tissues were fixed for 2h at 4°C in 4% paraformaldehyde in PBS. Fixed tongues were cut to obtain anterior two thirds to include fungiform (FP) and filiform (FILIF) papillae and the posterior third to include circumvallate (CV) and foliate (FOP) papillae. After fixation, all tissues were cryoprotected overnight with 30% sucrose in PBS and embedded in O.C.T. compound (Tissue-Tek, Sakura Finetek) for X-Gal staining or immunostaining as described previously [26,27]. Serial sagittal sections (FP), horizontal sections (CV, FOP, GG and TG) or coronal sections (SP) were cut at 10 μm for immunostaining [26,27].

X-gal staining

Tissue sections were incubated in X-gal solution for 16-18h except for lingual Gas1 and Gli3 tissues which were incubated for 4h and 48h, respectively. X-gal staining was followed by immunostaining of taste bud cells (K8) or epithelium (Ecad) in FP sections and ligand (SHH) in GG and TG sections.


Immunoreactions were performed as described previously [26,27]. Briefly, tongue or ganglion sections were air dried, rehydrated, blocked (in 10% normal donkey serum, 0.3% Triton-X in PBS-X), and incubated overnight at 4°C with primary antibodies. On the next day, slides were washed and incubated with appropriate secondary antibodies for 1–2 h at room temperature in the dark. Primary antibodies were goat anti-SHH (AF464, 0.1 μg/mL; R&D Systems); rat anti-keratin 8 (TROMA-1, 1:1,000; Developmental Studies Hybridoma Bank); goat anti–E-cadherin (AF748, 1:5,000; R&D Systems) and rabbit anti-RFP (600-401-379, 1:1,000; Rockland). For SHH in tongue sections, the heat-induced antigen-retrieval method [26] was used.


Tissue section images were acquired with a Nikon Eclipse 80i microscope and Nikon DS Ri2 camera system or an automated Keyence BZ-X810 microscope. Photomicrographs were adjusted for brightness and contrast in parallel across one figure and assembled with Adobe Photoshop.

Data analyses

We examined a minimum of 8 FP per tongue for each reporter line at each developmental stage. We also utilized 2–4 mouse tongues of the same postnatal age for analyzing FP, CVP and FOP, 2–3 SP, and GG and TG per line. X-gal expression was studied in serial sections, and the consistent presence of blue-colored product was considered as expression. We examined expression in the cell types depicted in the Fig 1A legend. The expression is categorized as ‘no’ if there is no blue product in that area, ‘partial’ if only few cells within the region are stained blue and ‘complete’ indicates that all cells are positive for X-gal staining. The expression patten was confirmed in replicates for all the different genes.


Expression pattern of Shh ligand and membrane receptors Ptch1, Gas1, Cdon and Hhip in the anterior tongue

In Fig 1A specific cell types in FP are illustrated. We compared expression patterns of HH pathway components across three developmental stages (early postnatal at P7, late postnatal at P28 and adult between 8–15 weeks of age). Shh ligand is present in FP taste buds, including taste bud basal cells, as observed with X-gal staining of the ShhlacZ reporter mouse model at P7, P28 and adult tongues (Fig 2A–2C) and align with previous studies using the SHH antibody detection method [2527]. We do not observe ligand expression in nerves with X-gal staining as reported with transgenic reporter models for Shh [26,4547].

Fig 2. Expression of HH pathway ligand and receptors in the anterior tongue from early postnatal to adult stages.

(A-O) X-gal staining using ShhlacZ/+, Ptch1lacZ/+, Gas1lacZ/+, CdonlacZ/+ and HhiplacZ/+ reporter mice at P7, P28 and adult stages. Shh ligand is expressed within taste buds at P7 (A), P28 (B) and adult tongue (C). Ptch1lacZ is expressed in the FP epithelium at all stages (D-F), but is predominant in the apical region (E, F arrows) from late postnatal to adulthood. In addition, Ptch1lacZ is expressed in filiform papillae (FILIF) epithelium (D-F, arrowheads) and tongue stromal cells. Gas1lacZ is expressed in taste buds at all stages (G-I), extensively in the stroma at P7 (G) and additionally in the epithelium from P28, albeit with less extensive stromal expression (H,I). CdonlacZ is observed in the entire lingual epithelium, within FP, it is concentrated in the lower half of the epithelium (J-L, arrows) and within FILIF, it appears not to overlap with Ptch1 locations (J-L, asterisks). Stromal CdonlacZ expression is also decreased from P7 to adult stages (J-L). The HH antagonist HhiplacZ is observed in FILIF at all the stages (M-O). HhiplacZ expression is also observed in the FP connective tissue core at P7 (M, arrow). Dotted lines outline the epithelium in all images. Scale bars (50μm) in A, B and C apply to respective column images.

The HH membrane receptor Ptch1 is expressed in both taste FP and non-taste FILIF from P7 through adult stages, as indicated by X-gal staining (Fig 2D–2F). However, the pattern in FP shows a shift between early and late postnatal stages. In FP at P7, Ptch1lacZ expression is observed in taste buds, perigemmal, apical extragemmal, basal and stromal cells (Fig 2D). At P28, there is no Ptch1lacZ expression within taste buds and expression in basal epithelial cells is limited to apical regions of FP, next to perigemmal cells (Fig 2E, arrow), while perigemmal, apical extragemmal and stromal cell expression are maintained (Fig 2E). This restricted basal cell pattern is also observed in the adult stage (Fig 2F, arrow). Upon investigating an earlier stage, P19, we again find a reduction in Ptch1lacZ expression in FP basal cells and loss of expression within taste buds (S1A Fig). In contrast, Ptch1lacZ expression in FILIF anterior epithelial face in cells of the basal and suprabasal layers is maintained from early postnatal stages to adulthood (Figs 2D–2F, arrowheads; S1B).

X-gal staining of Gas1lacZ mouse tongue revealed expression in FP taste buds from P7 to adult stages (Fig 2G–2I). While no other epithelial Gas1lacZ expression is present at P7 (Figs 2G and S1C, Ecad), expression emerges in the basal lingual epithelium on and after P21 (Figs 2H and 2I and S1D and S1E). In FP, Gas1lacZ expression is virtually absent in perigemmal and apical extragemmal cells, and the pattern is maintained through the adult stage (Fig 2G–2I). Further, we find extensive Gas1lacZ+ cells in the lingual stroma (Fig 2G) and muscles (S1C Fig) at or before P7. However, stromal expression declines while expression in muscle ceases on and after P21 (Figs 2H and 2I and S1D and S1E). We find that Gas1lacZ expression changed between P7 and P21, as indicated by a reduction in stromal cells, elimination in muscle cells and simultaneous emerging expression in lingual epithelial cells.

Another membrane HH co-receptor, CdonlacZ, is expressed in the entire lingual basal epithelium (but not in the apical half of FP wall) from P7 to the adult stage (Fig 2J–2L). This is unlike other HH receptors, Ptch1 and Gas1, which change their lingual epithelial expression from early postnatal stages to adulthood. Intriguingly, in FP, CdonlacZ expression is predominant in the basal lower half of FP (Fig 2J–2L, arrows) and in FILIF appears not to overlap with Ptch1 expression (Fig 2J–2L, asterisks). Although epithelial CdonlacZ expression in FP and FILIF is retained, its expression in overall tongue stromal cells showed reduction at eight weeks of age as compared to P7 or P28 (Figs 2J–2L and S1F and S1G). There is no Cdon lacZ expression in taste buds, perigemmal and apical extragemmal cells in any of the tongue developmental stages (Figs 2J–2L and S1F and S1G).

Similar to our recent study [27], HH antagonist HhiplacZ expression is observed in FILIF apical cells, consistently from P7 to adult stages (Fig 2M–2O). We do not observe HhiplacZ expression in taste buds, perigemmal and apical extragemmal cells (Fig 2M–2O). We observe expression of HhiplacZ in a few stromal cells within FP connective tissue core concentrating below the taste buds and in tongue stromal cells at P7. Stromal Hhip expression is noted at P12 (S1H Fig arrows) but not at P28 and adult stage (Figs 2N and 2O and S1I).

Overall, the data indicate that all transmembrane HH-receptors are expressed in the anterior tongue with partial to no co-expression, suggesting distinct and non-overlapping function in the development and maintenance of taste and non-taste organs. Intriguingly, we observed shifts in the unique expression patterns of Ptch1, Gas1 and Hhip in the early postnatal period primarily between P7-P21, suggesting a shift in function or a multi-functional role. Among the four HH membrane receptors studied here, only Gas1lacZ is consistently present in taste buds.

Transcription factor Gli1 expression changes during initial postnatal weeks, whereas Gli2 and Gli3 expressions remain stable in anterior tongue

As reported earlier [9,2527,48], Gli1lacZ expression is seen in FP basal, perigemmal, apical extragemmal and stromal cells and is maintained throughout postnatal and adult stages (Fig 3A–3C). However, we additionally observed Gli1lacZ+ cells in FILIF at P7 (Fig 3A, arrows), which are not seen after P28 (Fig 3B and 3C). To determine when Gli1 expression becomes restricted in the FP, we stained Gli1lacZ mouse tongues from a later developmental timepoint (P14). We found that 50% of FP lost lacZ expression in neighboring FILIF (S1J Fig). After P28, Gli1lacZ+ cells are observed only in FP (Fig 3B and 3C). Further, the expression is noted in basal and a few suprabasal cells at P7 (Fig 3A, arrowheads) but is restricted to FP basal cells only after P28 (Fig 3B and 3C).

Fig 3. HH pathway transcription factors expression in the anterior tongue from early postnatal to adult stages.

(A-I) X-gal staining of Gli1lacZ/+, Gli2lacZ/+and Gli3lacZ/+ reporter mice at the P7, P28 and adult stages. Gli1lacZ is expressed in FP basal, perigemmal, apical extragemmal and stromal cells (A-C). In addition, at P7, Gli1lacZ is observed in FILIF (A, arrows) and FP suprabasal epithelial layers (A, arrowheads). Gli2lacZ expression is observed in the entire lingual basal epithelium, suprabasal layers (D-F, arrows), perigemmal, apical extragemmal and stromal cells in (D-F). In contrast to Gli1lacZ and Gli2lacZ, Gli3lacZ is only present in lingual stromal cells (G-I). Dotted lines outline the epithelium in all images. Scale bars (50μm) in A,B and C apply to respective column images.

Gli2 reporter gene expression is observed in the entire lingual basal epithelium, perigemmal, apical extragemmal cells (including both FP and FILIF), and also in stromal cells from P7 to adult stages in Gli2lacZ mice (Fig 3D–3F). In addition, a few suprabasal epithelial cells of both FP and FILIF show Gli2lacZ expression (Fig 3D–3F, arrows). Stromal Gli2lacZ expression in the tongue is extensive as compared to Gli1lacZ expression throughout all stages (Fig 3D–3F). There is no overt change or shift in the expression pattern of Gli2lacZ in the adult tongue as compared to the early postnatal stages.

X-gal staining of Gli3lacZ mouse tongues revealed positive cells throughout the lingual stroma in both FP and FILIF, which remained unaltered from P7 to adult stages (Fig 3G–3I). Unlike other Gli transcription factors, Gli3 is not expressed in any of the epithelial (basal, perigemmal and apical extragemmal) cells. We found faint Gli3lacZ expression in a few taste buds in the adult stage after 48 hours of X-gal staining. A previous study in adult mice reported Gli3+ taste buds cells by in situ hybridization using digoxigenin-labeled Gli3 RNA probes [49].

The data suggest somewhat overlapping patterns of epithelial expression of Gli1 and Gli2 in FP and FILIF during initial postnatal stages. Similar Gli1 and Gli2 expression in FP are retained in late postnatal through adult stages. In contrast, Gli3 is observed only in stromal cells. Whether there is any overlap between Gli stromal expressions was not studied.

Mapping Shh ligand and membrane receptors Ptch1, Gas1, Cdon and Hhip expression in posterior circumvallate and foliate papillae at early postnatal and adult stages

Illustrations for CVP and FOP structure and cell types in the horizontal plane are shown in Fig 1A. Given the continuous postnatal development of posterior CVP and FOP taste buds [10], we studied the initial postnatal week with limited taste bud formation and mature adult stages with a full complement of taste buds to analyze HH pathway component expression (Figs 4 and 5). Immunostaining with SHH antibody or X-gal staining of the ShhlacZ reporter mouse model at P7 showed ligand expression in CVP and FOP taste buds (Figs 4A and 4C and K8+; S1K). To visualize Shh expression in nerves, we used tissues from ShhCreERT2; R26RFP animals to determine whether nerves are labeled in adult CVP and FOP. After 30 days of tamoxifen induction Shh expression is observed in nerve fibers surrounding the CVP walls and all FOP ridges in the connective tissue core (Fig 4B and 4D, arrows) in addition to taste buds. Previous work using the ShhCreER/+; R26mTmG model also showed Shh+ fibers within CVP taste buds [47]. CVP and the posterior-most folds of the FOP are innervated by glossopharyngeal nerves (Fig 1A), which have soma in the petrosal ganglion. Shh expression in the petrosal ganglion is not yet studied.

Fig 4. Expression of HH pathway ligand and receptors in posterior tongue circumvallate and foliate papillae from early postnatal and adult stages.

(A-D) SHH ligand expression within CVP and FOP taste buds (labelled with K8, green, A) as demonstrated by either antibody staining or X-gal staining in ShhlacZ/+ reporter mouse at P7 (A,C) and RFP staining in ShhCreERT2;R26RFP reporter mouse after 30 days of tamoxifen treatment at adult stage (B,D).(E-T) X-gal staining using ShhlacZ/+, Ptch1lacZ/+, Gas1lacZ/+, CdonlacZ/+ and HhiplacZ/+ reporter mice at P7 and adult stages. In both stages, Ptch1lacZ is expressed in basal, perigemmal, apical extragemmal and stromal cells of CVP (E, F) and FOP (G,H). A few Ptch1lacZ punctae are seen within taste buds of CVP at both stages (E, F, arrows) and FOP at P7 (G, arrow). Gas1lacZ is present in taste buds and stromal cells at P7 and in adult CVP (I, J) and FOP (K, L). Stromal expression of CdonlacZ is observed in CVP (M,N) and FOP (O,P), but the expression declines at adult stages (N,P) as compared to P7 (M,O). Extensive HhiplacZ CVP and FOP stromal expression is observed at P7 (Q,S), which is virtually eliminated by the adult stage (R,T), and few punctate lacZ+ cells are observed in CVP stroma at adult stage (R, arrow). Black or white dotted lines outline the epithelium and yellow dotted lines outline the taste buds apical surface in all the images. Scale bars are 50μm.

For Ptch1lacZ expression, we investigated CVP and FOP sagittal sections at P7 and horizontal sections at the adult stage (Fig 4E–4H). We observed Ptch1lacZ+ cells in CVP and FOP basal, perigemmal, apical extragemmal and stromal cells at both stages (Fig 4E–4H). Magnification of the CVP wall shows few punctate Ptch1lacZ+ cells within taste buds at either P7 or adult stages (Fig 4E and 4F, arrows). Little Ptch1lacZ expression is seen within FOP taste buds at P7 (Fig 4G, arrow).

X-gal staining of horizontal sections of the Gas1lacZ mouse posterior tongue at P7 and adult revealed expression in CVP and FOP taste buds (Fig 4I–4L). There is extensive Gas1lacZ expression in CVP stroma, but stromal cells of CVP adjacent to papilla walls had few Gas1lacZ+ cells as compared to overall Gas1lacZ CVP stromal expression at both the stages (Fig 4I and 4J). Gas1lacZ is expressed throughout FOP stroma (Fig 4K and 4L).

Contrasting with Gas1, CdonlacZ expression is not seen in either CVP or FOP taste buds (Fig 4M–4P). The CdonlacZ+ stromal cell expression pattern at P7 in the CVP connective tissue core is comparable to the Gas1lacZ arrangement, with few positive cells surrounding papilla walls whereas the expression is prominent in the rest of the stromal cells distant to papilla walls (Fig 4M). We observed CdonlacZ+ cells throughout FOP stromal cells at P7 (Fig 4O). In both CVP and FOP at adult stages (Fig 4N and 4P), the expression declined as compared to P7 (Fig 4M and 4O). It is possible that the remaining punctate X-gal staining represents residual β-gal enzyme rather than active gene expression [50]. Unlike the anterior FP expression pattern (Fig 1J–1L), we do not observe CdonlacZ expression in CVP or FOP basal cells (Fig 4M–4P).

In contrast to Gas1lacZ and CdonlacZ expression patterns in the connective tissue core, HhiplacZ is extensively expressed in CVP and FOP stromal cells neighboring the papilla walls at P7 (Fig 4Q and 4S). Further, HhiplacZ expression is limited to a few stromal cells in adult CVP and virtually absent in the adult FOP connective tissue core, as indicated in horizontal sections (Fig 4R, arrow; 4T). The stromal CVP and FOP Hhip expression at an early developmental stage (Fig 4Q and 4S) is coincident with expression in FP stroma (S1H Fig).

Overall, the expression patterns of Shh ligand and HH membrane receptors Ptch1, Gas1, and Cdon remain unchanged during the postnatal development of CVP and FOP. Only Cdon and Hhip expression changed in the CVP and FOP connective tissue core at 8 weeks of age as compared to the first postnatal week. Interestingly, basal epithelial expression of Gas1 and Cdon, as seen in anterior tongue, is not observed in the posterior CVP and FOP at any given developmental stage(s). The data indicate the requirement of different HH receptors between anterior and posterior tongue epithelial development.

Gli transcription factor expression in the posterior circumvallate and foliate papillae remain the same throughout postnatal development, while Gli3 is additionally expressed in papilla taste buds in adults

In both P7 and adult tongues, we observed Gli1lacZ expression in CVP and FOP basal, perigemmal, apical extragemmal and stromal cells (Fig 5A–5D). Similar to Gli1lacZ, Gli2lacZ+ cells are present in CVP and FOP basal, perigemmal, apical extragemmal and stromal cells throughout postnatal and adult stages (Fig 5E–5H). There is no Gli1lacZ or Gli2lacZ expression within taste buds of CVP or FOP (Fig 5A–5H).

Fig 5. Expression pattern of HH pathway transcription factors in circumvallate and foliate papillae from early postnatal and adult stages.

(A-L) X-gal staining of Gli1lacZ/+, Gli2lacZ/+and Gli3lacZ/+ reporter mice at P7 and adult stages. Gli1lacZ (A-D) and Gli2lacZ (E-H) are expressed in basal, perigemmal, apical extragemmal and stromal cells of CVP (A,B,E,F) and FOP (C,D,G,H) at both the P7 and adult stages. Gli3lacZ is present in CVP and FOP stromal cells (I-L) at P7 (sagittal section) and adult tongue (horizontal section) and additionally in taste buds in adult CVP and FOP taste buds (J,L arrows). Black dotted lines outline the epithelium and yellow dotted lines outline the taste buds apical surface in all the images. Scale bars (50μm) in B,C and D apply to respective column images.

X-gal staining of P7 sagittal tongue sections revealed Gli3lacZ+ cells in CVP and FOP stroma (Fig 5I and 5K). Gli3lacZ expression in CVP and FOP stroma surrounding papilla walls remains similar in adult horizontal tongue sections (Fig 5J and 5L) as compared to P7 (Fig 5I–5K). In addition, there are positive cells in CVP and FOP taste buds in mice older than 8 weeks (Fig 5J and 5L, arrows).

The data indicate that Gli transcription factor expression in CVP and FOP mostly remains the same from early postnatal weeks through adult stages, and the location patterns are similar to each other and to the anterior tongue.

HH signaling component expression in adult soft palate is similar to fungiform papilla

We investigated the soft palate (SP) with taste organs in the oral cavity that harbor taste buds distributed in the epithelium, not enclosed in papillae. We limited our investigations to the adult stage. X-gal staining of the ShhlacZ mouse SP reveals expression in taste buds (S1L Fig) consistent with previous findings [51]. Further, with the 14-day tamoxifen-induced ShhCreERT2;R26RFP mouse model, Shh is observed in both taste buds and nerves (Fig 6A). Different transmembrane HH receptors have distinct expression: Ptch1lacZ in SP perigemmal and stromal cells with some punctate expression within taste buds cells (Fig 6B); Gas1lacZ in SP basal epithelial, stromal and taste buds cells (Fig 6C); CdonlacZ in the SP basal epithelium, mainly in the lower half (Fig 6D). We did not observe HhiplacZ expression in SP (Fig 6E) as reported earlier [27]. All HH transcription factors Gli1lacZ, Gli2lacZ and Gli3lacZ are expressed within SP stromal cells (Fig 6F–6H). Gli1lacZ and Gli2lacZ are additionally expressed in SP basal epithelial, perigemmal and apical extragemmal cells (Fig 6F and 6G). Our HH component expression data reveal active HH signaling in SP, similar to FP.

Fig 6. Expression of HH pathway components in adult mouse soft palate.

(A) RFP staining in ShhCreERT2;R26RFP reporter mouse after 14 days of tamoxifen treatment reveals Shh expression in the soft palate (SP) taste buds and nerves (A, arrow). (B) Ptch1lacZ is expressed in perigemmal and stromal cells along with punctate expression within taste buds. (C) Gas1lacZ is present within the taste buds, epithelium and stroma. (D) CdonlacZ expression is seen at the SP epithelium rete ridges (D, arrows). (E) HhiplacZ is not expressed in the SP. (F-G) Both Gli1lacZ (F) and Gli2lacZ (G) are present in SP basal, perigemmal, apical extragemmal and stromal cells. (H) Gli3lacZ is exclusively present in the stromal cells. Black or white dotted lines outline the epithelium and orange dotted lines outline the taste buds in all the images. Scale bar (50μm) in A applies to all images.

SHH ligand, membrane receptor Ptch1, and transcription factors Gli1 and Gli2 are the only HH pathway components expressed in adult geniculate and trigeminal ganglia

Geniculate ganglion (GG) and trigeminal ganglion (TG) contain cell bodies of chorda tympani and lingual nerves, respectively, that innervate the anterior tongue [52]. Previous studies with reporter mice indicate Shh ligand expression in the adult mouse GG [26,45,47] and TG [26]. Here, we have used a SHH antibody and reproduced the finding of SHH+ soma in both GG (Fig 7B, 7F, 7J and 7N) and TG (Fig 7D, 7H, 7L and 7P). Among all the HH co-receptors, Ptch1 is the only one expressed in both GG and TG (Fig 7A–7D). We analyzed Ptch1 expression qualitatively based on location within the neuron and outside the neuron, touching either the cell body or the nerve bundles exiting the ganglion. Our data indicate that the Ptch1lacZ+ cells either overlap (arrows) or are adjacent to (arrowheads) SHH+ neuronal cells in adult GG and TG (Fig 7B and 7D). There is no expression of Gas1lacZ (Fig 7E–7H), CdonlacZ (Fig 7I–7L) or HhiplacZ (Fig 7M–7P) in either GG or TG.

Fig 7. Expression pattern of HH ligand and receptors in adult mouse geniculate and trigeminal ganglia.

(A-P) X-gal staining in Ptch1lacZ/+, Gas1lacZ/+, CdonlacZ/+ and HhiplacZ/+ reporter mice. SHH antibody staining indicates SHH+ (red) cell bodies in GG (B, F, J, N) and TG (D, H, L, P). Ptch1lacZ expression is observed both in GG (A, B) and TG (C, D) and overlaps (arrows) or neighbors (arrowheads) SHH+ cells (B, D red). Yellow lines demarcate nerve bundles exiting the ganglion. None of the other HH-co-receptors, Gas1lacZ (E-H), CdonlacZ (I-L) and HhiplacZ (M-P) is expressed in either GG or TG. Scale bars (50μm) in A and C apply to all Geniculate Ganglion and Trigeminal Ganglion images, respectively.

Among the transcription factors, Gli1lacZ and Gli2lacZ, but not Gli3lacZ are observed in both GG and TG (Fig 8A–8L). Further, Gli1lacZ+ and Gli2lacZ+ cells are either next to SHH+ cells (arrowhead) or within the nerve bundles exiting the ganglion (asterisk) (Fig 8B, 8D, 8F and 8H). Importantly, in our observations not all SHH+ neurons have adjacent Gli1lacZ+ and Gli2lacZ+ cells suggesting specificity towards a neuron cell type for a particular function.

Fig 8. Expression pattern of HH transcription factors in adult mouse geniculate and trigeminal ganglia.

(A-L) X-gal staining in Gli1lacZ/+, Gli2lacZ/+ and Gli3lacZ/+ reporter mice. SHH antibody staining indicates SHH+ (red) cell bodies in GG (B, F, J) and TG (D,H,L). Both Gli1lacZ (A-D) and Gli2lacZ (E-H) expressions are observed in GG (A,B,E,F) and TG (C,D,G,H) with (arrowheads) and without neighboring SHH ligand (B,D,F,H red). Additionally, both are expressed within the nerve bundles exiting the ganglion (B, D, F, H, asterisks). Yellow lines demarcate nerve bundles exiting the ganglion. In contrast, the transcription factor Gli3lacZ is not expressed in either GG or TG (I-L). Scale bars (50μm) in A and C apply to all Geniculate Ganglion and Trigeminal Ganglion images, respectively.

Overall, the presence of SHH, Ptch1, Gli1 and Gli2 indicates a requirement for active HH signaling in GG and TG while the absence of Gas1, Cdon, Hhip and Gli3 implies that there is no HH antagonism in either ganglion.


Here we have analyzed the expression of HH pathway ligand, membrane receptors and antagonist, and the GLI transcription factors in the gustatory system during development and/or homeostasis. We have examined HH pathway component expressions across six distinct tissues, FP, CVP, FOP, SP, GG, and TG (Table 1). Our data unveil unique expression patterns in the lingual epithelium, stroma and muscles. The identified shifts in gene expressions during late postnatal stages match with the conclusion of anterior tongue morphogenesis by postnatal day 21 [10,15,16,18]. Further, the HH pathway receptors, Gas1 and Cdon, which remain unexplored in the tongue, have extensive yet dynamic lingual epithelium, stromal and muscular expressions.

Table 1. Summary of HH pathway component expressions in gustatory system.

Shh ligand is present in taste buds and nerves of tongue and soft palate

While previous studies have demonstrated that SHH is expressed in FP taste bud cells and nerves [9,26,45,47,53], data on other taste organs are lacking and focused mainly on the adult stage. Here we show that Shh ligand is consistently expressed within the taste buds and in the nerves entering taste buds of all tongue papillae and soft palate (Table 1). Whereas nerve-derived HH ligands participate in touch dome [54,55] and hair follicle [56] maintenance, similar effects in postnatal or adult FP taste cell differentiation were not observed [8,45]. The roles of neural SHH on posterior tongue and SP taste buds have not yet been evaluated.

Recently, it has been shown that misexpression of Shh in lingual epithelium induces ectopic taste bud formation in FILIF and the number of ectopic taste buds is 3-fold higher when overexpression occurs at P14 as compared to P1 [8]. In this study, our data indicate that expression of the HH receptor Gas1 is induced in FILIF at later postnatal stages, which may explain the temporal rise in ectopic taste buds reported in the previous study.

HH signaling is active in anterior taste and non-taste papilla epithelium during early postnatal tongue development

HH signaling, as indicated by the target gene, Gli1, in adult tongue is restricted to taste organs [27]. For postnatal stages, a previous study utilized in situ staining and showed Gli1 expression in FP only [9]. However, in situ staining might have limitations for detecting lower gene expression [57]. Thus, we utilized Gli1 reporter mouse models and confirmed Gli1+ cells in FP and intriguingly revealed additional expression in FILIF in the initial postnatal weeks. This expression, however, disappears from FILIF by P21 and beyond. Notably, FILIF grow rapidly in the first postnatal week and steadily thereafter until P21, when they reach full maturation [58]. Given that HH/SMO/GLI inhibition at the adult stage did not alter FILIF structure and pattern [27], we propose that HH signaling regulates FILIF maturation and after the conclusion of postnatal morphogenesis by P21, FILIF become HH-independent.

HH co-receptors are expressed in tongue taste and non-taste papilla epithelium

We recently reported the expression of the HH receptor Ptch1 in FP and anterior epithelial face of FILIF and suggested dual roles for the receptor: activator in FP and antagonist in FILIF [27]. In addition to PTCH1, there are additional co-receptors, GAS1, BOC and CDON, that help create the SHH gradient during organ development [34,35,5963] but remain unexplored in the tongue. Here, we have determined both distinct and overlapping spatiotemporal expression patterns of Gas1 and Cdon in the postnatal lingual epithelium, including both FP and FILIF.

Unlike other receptors, extensive Gas1 expression within the taste bud of all three taste papillae suggests Gas1-mediated SHH signaling for taste cell differentiation. Gas1 expression is seen in the basal lingual epithelium after P21. HH signaling becomes restricted to FP with concurrent expression of Gas1 in the postnatal lingual epithelium. Initial studies claimed Gas1 as an antagonist of HH signaling [59,64] while subsequent studies showed positive regulation by Gas1 [19,20,63]. As Gas1 function can be ligand-independent [65], we propose that Gas1 may have distinct functions in taste buds, FP and FILIF cell contexts depending on ligand availability.

Cdon expression remains the same in the anterior lingual epithelium throughout postnatal stages, irrespective of the changes in Gli1 and Gas1 expression. Cdon expression, which co-occurs with Gli1 in the early postnatal stage and overlaps with Gas1 in the late postnatal stage, suggests its potential role as a multifunctional regulator of HH signaling during tongue maturation. This role may involve promoting HH signaling in FILIF during early stages and later inhibiting it in collaboration with Gas1. Interestingly, this cooperation between Gas1 and Cdon was not observed during limb development [19], even though they are expressed in overlapping domains in the limb [62,63]. Our data further suggest that Cdon may not overlap with Ptch1 suggesting that SHH employs Cdon, Gas1 and Ptch1 in a context-dependent manner during early and late postnatal stages of tongue development.

HH co-receptors are differentially expressed in tongue stroma

While previous work has documented Ptch1 and Gli expression in adult lingual stromal cells [9,25,26,48,49], here we identified spatiotemporal expression patterns of two HH co-receptors, Gas1 and Cdon and the antagonist Hhip in postnatal lingual stroma (Table 1). HH signaling-mediated epithelial–mesenchymal interactions are critical in the development of many tissues, including tooth [66] and palate [67]. In embryonic tongue, stromal cells are involved in the transmission of information from epithelial cells to myogenic progenitor cells for muscle maturation [68,69]; at P14, stromal cells can generate a few taste bud cells [70], but whether they are HH-responsive is unknown. In our epithelial HH/GLI/SMO repression in adult mouse models, stromal cells were retained but could not prevent taste cell loss [25,26], however, this does not rule out a postnatal stromal contribution to FP taste buds.

In limb bud, Cdon+ HH responding cells extend long specialized filopodia relaying activation of the pathway at a distance from the cell soma [21]. Similarly, we observed vimentin+ stromal cells with filopodial extensions that contact the basal lamina [25], but whether they are Cdon+ is not known. In this study, we observed a decline in HH co-receptor expressions in all tongue papillae stroma at later postnatal stages. Further, the opposing expression patterns of Hhip with Gas1 and Cdon in CVP and FOP suggest a discrete population of stromal cells, neighboring and distant to the papillae walls. We hypothesize that stromal HH signaling might provide structural support and regulate the development of lingual epithelium and muscle in the early postnatal stages.

Gas1 is the only HH pathway component identified in lingual muscles at early postnatal stage

While HH signaling guides embryonic muscle development [71,72], we found no Shh or HH-responding Gli1 expression in postnatal lingual muscles. This argues against direct HH control of tongue muscles, consistent with other studies suggesting stroma-mediated regulation of muscle development [68,69]. Here, we identified Gas1 expression in postnatal lingual muscles (Figs 2G and S1C) during their crucial maturation phase [73,74]. However, a role for Gas1 in the postnatal tongue is not yet known. Our recent studies revealed that global Gas1 deletion did not alter embryonic myoblast migration but caused defects in muscle differentiation leading to altered muscle arrangement [75]. We propose that in addition to its embryonic effect Gas1 can also have cell-autonomous effects on lingual muscles in early postnatal stages.

Distinct expression pattern of HH signaling components in anterior and posterior tongue at adult stage

Building on our previous work exploring anterior-posterior tongue regulatory mechanisms [52], here we reveal similarities and differences in HH signaling activity within adult taste papillae (Fig 9). Shh ligand and the HH receptor Gas1 exhibit similar expression patterns across all three papilla taste buds, suggesting a conserved homeostatic mechanism. On the other hand, Gli3 is observed predominantly in posterior papilla taste buds (Fig 9). This contrasts with Gas1 and Cdon, which are expressed in anterior tongue papilla epithelium (Fig 9), potentially contributing to the restriction of HH signaling to adult FP.

Fig 9. Expression patterns of HH signaling components in the adult FP, CVP, FOP and soft palate.

Building from Fig 1A, HH signaling components are located in FP, CVP, FOP and soft palate. SHH is predominant in taste bud basal cells. Gas1 is present in all papillae taste bud, whereas Gli3 is also expressed in the posterior tongue papillae taste bud. Hhip and Ptch1 are present in anterior epithelial face of FILIF. Gli1+, Ptch1+ and Gli2+ cells are observed in lingual basal, perigemmal, apical extragemmal and stromal cells of all the papillae. However, Ptch1 is primarily present in the upper apical half of FP. On the other hand, HH co-receptors Gas1 and Cdon are concentrated in lower half of FP wall and entire lingual epithelium. Gas1 and Gli3 are also expressed in the stromal cells of all the papillae and soft palate. The expression pattern of HH signaling components in the soft palate is similar to that observed in FP.

Ptch1, Gli1 and Gli2 are present in both anterior and posterior lingual basal, perigemmal and apical extragemmal cells (Fig 9). While Ptch1+ cells from FP basal cells become restricted to the apical half of FP (anterior tongue), apparently all basal cells in posterior tongue papillae retain Ptch1 expression. A similar decline in Ptch1 expression is also observed in the dental inner enamel epithelium to maintain SHH-responsive quiescent adult stem cells [76]. Our previous HH pathway inhibition studies showed a reduction in cell proliferation [25,26,28] and elimination of Ptch1 [27], in apically lying basal cells, implying that Ptch1 controls FP apical cell proliferation. While the basal half of FP retained cell proliferation, it could not prevent taste bud loss [25,26,28], which further suggests FP taste bud stem cell maintenance by Ptch1+ cells. The data highlight heterogeneous cell populations in the FP basal epithelium, which demarcate boundaries between the apical and basal epithelium (Fig 9). Further, the HH antagonist Hhip is specific to non-taste FILIF and is not observed in the taste papilla epithelium (Fig 9).

Stromal expression of HH receptors and GLI transcription factors is observed but whether they are co-expressed in the same subset of stromal cells remains to be investigated. We propose stromal diversity in the context of HH signaling components to create a distinct niche, as described previously [77].

HH signaling in other oral tissue at adult stage

In addition to FP, taste buds are also housed in the soft palate (SP) epithelium without being enclosed in a papilla (Fig 9, soft palate). The maturation of SP taste buds is almost complete by P7 and is much faster than that of the taste buds of tongue papillae (after P21) [10]. Upon analyzing SP taste buds in adult mice, we found expression patterns similar to FP (Fig 9). Shh and Gas1 are co-expressed in taste buds. The expression patterns of the HH receptors Ptch1 and Gas1, and all three Gli transcription factors in mesenchyme corroborate a previous study conducted between embryonic days 13 and 14.5 [78]. In contrast to previous embryonic SP data [78], we observe epithelial expression of Ptch1, Gas1, Gli1 and Gli2 but not expression of Hhip in the adult SP. Whether these differences reflect a change in HH component expression or differences in technical approaches [78] remains to be investigated. Although Cdon was not studied previously, here we observed an expression pattern in the epithelium similar to Gas1.

We show that HH-responding Gli1+ cells are present in the SP epithelium, which give rise to taste buds [79]. When we treated adult rats with the HH pathway inhibition drug sonidegib, we observed that taste buds were reduced to half their normal numbers and the remaining taste buds were atypical [28] suggesting that HH signaling regulates adult SP taste buds. Embryonic SP epithelial-mesenchymal interactions, mediated by HH signaling, control palatal outgrowth [68,8082]. Beyond embryonic HH mesenchymal signaling, in adult SP, our expression studies suggest paracrine signaling regulation from Shh+ taste buds and nerves to HH-responding palatal perigemmal, basal and stromal cells for tissue homeostasis.

HH signaling in ganglia associated with gustatory system during adulthood

Multimodal chorda tympani, greater superficial petrosal and somatosensory lingual nerves receive afferents from the geniculate (GG) and trigeminal ganglion (TG) soma, respectively [83]. SHH is expressed in all GG and TG neurons [26,45]. Therefore, we see Shh+ nerves within the taste buds of FP that receive chorda tympani nerve fibers and SP that receive greater superficial petrosal nerve fibers. Shh expression in the lingual nerve innervating FP walls and FILIF connective tissue core is not clearly demonstrated even though TG neurons express SHH [28,46,47]. It is unclear whether this is a detection issue or reflects previously unappreciated neuronal heterogeneity.

Here, we reveal the expression patterns of other members of HH signaling (Table 1). Compared to the tongue or the soft palate, the GG and TG might utilize only positive regulators of HH signaling. While our findings indicate HH signaling likely regulates GG and TG, its precise impact on neuronal function and development needs further investigation.

In conclusion, our comparative analyses of unexplored HH pathway co-receptors and partially explored GLI transcription factors in anterior and posterior tongue, soft palate and ganglia, in epithelium, stroma or muscles reveals heterogeneity in their expression patterns. This work lays a foundation for understanding the intricate roles of HH signaling in distinct organs of the gustatory system. Future functional studies, guided by these expression maps, will dissect the regulatory mechanisms by which HH signaling components orchestrate tissue-specific development and function within the gustatory system.

Supporting information

S1 Fig. Expression pattern of HH signaling components in the anterior and posterior tongue and soft palate.

(A, B) X-gal staining in Ptch1lacZ/+ reporter mouse indicates a reduction in Ptch1+ FP basal cells and taste bud cells at P19 (A) and expression in FILIF anterior epithelial face at P7 (B). (C-E) X-gal staining in Gas1lacZ/+ reporter mouse at P4, P21 and adult stages suggests that while lingual muscles express Gas1lacZ at P4, Gas1+ muscle cells are not observed at P21 and adult tongues. Ecad antibody co-staining confirms absence of Gas1lacZ expression in lingual epithelium at P4 (C). Epithelial Gas1lacZ expression is observed at P21 (D) and maintained through the adult stage (E). (F,G) X-gal staining in CdonlacZ/+ reporter mouse reveals stromal expression at P8 (F), which gets downregulated at adult stage (G). K8 antibody co-staining confirms no Cdon lacZ expression in taste bud (F). (H,I) X-gal staining in HhiplacZ/+ reporter mouse shows stromal HhiplacZ expression at P12 in tongue (H) and below the taste bud (H, inset, arrow) but not at P30 (I). (J) X-gal staining in P14 Gli1lacZ/+ reporter mouse indicates reduced lacZ expression in FILIF (arrow). (K, L) X-gal staining in ShhlacZ/+ reporter mouse shows expression in the taste buds of CVP at P7 (K) and soft palate at adult stage (L). Black dotted lines outline the epithelium. Scale bars are 50μm.



We thank Dr. Chen-Ming Fan for sharing the Gas1lacZ mouse model with AK.


  1. 1. Hosley MA, Oakley B. Postnatal development of the vallate papilla and taste buds in rats. Anat Rec. 1987;218(2):216–22. pmid:3619089
  2. 2. Miller IJ Jr., Smith DV. Proliferation of taste buds in the foliate and vallate papillae of postnatal hamsters. Growth Dev Aging. 1988;52(3):123–31. pmid:3253244
  3. 3. Mistretta C. Smell and taste in health and disease. Raven Press, New York; 1991.
  4. 4. Mistretta CM, Bradley RM. Developmental changes in taste responses from glossopharyngeal nerve in sheep and comparisons with chorda tympani responses. Brain Res. 1983;313(1):107–17. pmid:6661660
  5. 5. Hall JM, Hooper JE, Finger TE. Expression of sonic hedgehog, patched, and Gli1 in developing taste papillae of the mouse. J Comp Neurol. 1999;406(2):143–55. pmid:10096602
  6. 6. Mistretta CM, Goosens KA, Farinas I, Reichardt LF. Alterations in size, number, and morphology of gustatory papillae and taste buds in BDNF null mutant mice demonstrate neural dependence of developing taste organs. J Comp Neurol. 1999;409(1):13–24.PMC2710125. pmid:10363708
  7. 7. Mistretta CM, Liu HX. Development of fungiform papillae: patterned lingual gustatory organs. Arch Histol Cytol. 2006;69(4):199–208. pmid:17287575
  8. 8. Golden EJ, Larson ED, Shechtman LA, Trahan GD, Gaillard D, Fellin TJ, et al. Onset of taste bud cell renewal starts at birth and coincides with a shift in SHH function. Elife. 2021;10.PMC8172241. pmid:34009125
  9. 9. Liu HX, Ermilov A, Grachtchouk M, Li L, Gumucio DL, Dlugosz AA, et al. Multiple Shh Signaling Centers Participate in Fungiform Papilla and Taste Bud Formation and Maintenance. Dev Biol. 2013;382(1):82–97.PMC3968530. pmid:23916850
  10. 10. Zhang GH, Zhang HY, Deng SP, Qin YM, Wang TH. Quantitative study of taste bud distribution within the oral cavity of the postnatal mouse. Arch Oral Biol. 2008;53(6):583–9. pmid:18294610
  11. 11. Oakley B. Reformation of taste buds by crossed sensory nerves in the rat’s tongue. Acta Physiol Scand. 1970;79(1):88–94. pmid:5464393
  12. 12. Oakley B, LaBelle DE, Riley RA, Wilson K, Wu LH. The rate and locus of development of rat vallate taste buds. Brain Res Dev Brain Res. 1991;58(2):215–21. pmid:2029765
  13. 13. Pritchard TC. The primate gustatory system. Smell and taste in health and disease. 1991:109–25.
  14. 14. Zhang GH, Chen ML, Liu SS, Zhan YH, Quan Y, Qin YM, et al. Facilitation of the development of fungiform taste buds by early intraoral acesulfame-K stimulation to mice. J Neural Transm (Vienna). 2010;117(11):1261–4. pmid:20838827
  15. 15. Zhang GH, Deng SP, Li LL, Li HT. Developmental change of alpha-gustducin expression in the mouse fungiform papilla. Anat Embryol (Berl). 2006;211(6):625–30. pmid:16933139
  16. 16. Harada S, Yamaguchi K, Kanemaru N, Kasahara Y. Maturation of taste buds on the soft palate of the postnatal rat. Physiol Behav. 2000;68(3):333–9. pmid:10716542
  17. 17. Huang T, Ma L, Krimm RF. Postnatal reduction of BDNF regulates the developmental remodeling of taste bud innervation. Dev Biol. 2015;405(2):225–36.PMC4572574. pmid:26164656
  18. 18. Ohtubo Y, Iwamoto M, Yoshii K. Subtype-dependent postnatal development of taste receptor cells in mouse fungiform taste buds. Eur J Neurosci. 2012;35(11):1661–71. pmid:22462540
  19. 19. Allen BL, Tenzen T, McMahon AP. The Hedgehog-binding proteins Gas1 and Cdo cooperate to positively regulate Shh signaling during mouse development. Genes Dev. 2007;21(10):1244–57.PMC1865495. pmid:17504941
  20. 20. Martinelli DC, Fan CM. Gas1 extends the range of Hedgehog action by facilitating its signaling. Genes Dev. 2007;21(10):1231–43.PMC1865494. pmid:17504940
  21. 21. Sanders TA, Llagostera E, Barna M. Specialized filopodia direct long-range transport of SHH during vertebrate tissue patterning. Nature. 2013;497(7451):628–32.PMC4197975. pmid:23624372
  22. 22. Allard BA, Wang W, Pottorf TS, Mumtaz H, Jack BM, Wang HH, et al. Thm2 interacts with paralog, Thm1, and sensitizes to Hedgehog signaling in postnatal skeletogenesis. Cell Mol Life Sci. 2021;78(7):3743–62.PMC9278483. pmid:33683377
  23. 23. Yie TA, Loomis CA, Nowatzky J, Khodadadi-Jamayran A, Lin Z, Cammer M, et al. Hedgehog and PDGF Signaling Intersect During Postnatal Lung Development. Am J Respir Cell Mol Biol. 2023.
  24. 24. Petrova R, Joyner AL. Roles for Hedgehog Signaling in Adult Organ Homeostasis and Repair. Development. 2014;141(18):3445–57.PMC4197719. pmid:25183867
  25. 25. Ermilov AN, Kumari A, Li L, Joiner AM, Grachtchouk MA, Allen BL, et al. Maintenance of Taste Organs is Strictly Dependent on Epithelial Hedgehog/GLI Signaling. PLoS Genet. 2016;12(11):e1006442.PMC5125561. pmid:27893742
  26. 26. Kumari A, Ermilov AN, Grachtchouk M, Dlugosz AA, Allen BL, Bradley RM, et al. Recovery of Taste Organs and Sensory Function after Severe Loss from Hedgehog/Smoothened Inhibition with Cancer Drug Sonidegib. Proc Natl Acad Sci U S A. 2017;114(48):E10369–E78.PMC5715770. pmid:29133390
  27. 27. Kumari A, Li L, Ermilov AN, Franks NE, Dlugosz AA, Allen BL, et al. Unique lingual expression of the Hedgehog pathway antagonist Hedgehog-interacting protein in filiform papillae during homeostasis and ectopic expression in fungiform papillae during Hedgehog signaling inhibition. Dev Dyn. 2022;251(7):1175–95. pmid:35048440
  28. 28. Kumari A, Yokota Y, Li L, Bradley RM, Mistretta CM. Species generalization and differences in Hedgehog pathway regulation of fungiform and circumvallate papilla taste function and somatosensation demonstrated with sonidegib. Sci Rep. 2018;8(1):16150.PMC6212413. pmid:30385780
  29. 29. Liu HX, Maccallum DK, Edwards C, Gaffield W, Mistretta CM. Sonic hedgehog exerts distinct, stage-specific effects on tongue and taste papilla development. Dev Biol. 2004;276(2):280–300. pmid:15581865
  30. 30. Mistretta CM, Liu HX, Gaffield W, MacCallum DK. Cyclopamine and jervine in embryonic rat tongue cultures demonstrate a role for Shh signaling in taste papilla development and patterning: fungiform papillae double in number and form in novel locations in dorsal lingual epithelium. Dev Biol. 2003;254(1):1–18. pmid:12606278
  31. 31. Briscoe J, Therond PP. The Mechanisms of Hedgehog Signalling and Its Roles in Development and Disease. Nat Rev Mol Cell Biol. 2013;14(7):416–29 pmid:23719536
  32. 32. Stone DM, Hynes M, Armanini M, Swanson TA, Gu Q, Johnson RL, et al. The tumour-suppressor gene patched encodes a candidate receptor for Sonic hedgehog. Nature. 1996;384(6605):129–34. pmid:8906787
  33. 33. Taipale J, Cooper MK, Maiti T, Beachy PA. Patched acts catalytically to suppress the activity of Smoothened. Nature. 2002;418(6900):892–7. pmid:12192414
  34. 34. Allen BL, Song JY, Izzi L, Althaus IW, Kang J-S, Charron F, et al. Overlapping roles and collective requirement for the coreceptors GAS1, CDO, and BOC in SHH pathway function. Dev Cell. 2011;20(6):775–87. pmid:21664576
  35. 35. Holtz AM, Peterson KA, Nishi Y, Morin S, Song JY, Charron F, et al. Essential role for ligand-dependent feedback antagonism of vertebrate hedgehog signaling by PTCH1, PTCH2 and HHIP1 during neural patterning. Development. 2013;140(16):3423–34.PMC3737722. pmid:23900540
  36. 36. Jeong J, McMahon AP. Growth and pattern of the mammalian neural tube are governed by partially overlapping feedback activities of the hedgehog antagonists patched 1 and Hhip1. Development. 2005;132(1):143–54. pmid:15576403
  37. 37. Hui CC, Angers S. Gli proteins in development and disease. Annu Rev Cell Dev Biol. 2011;27:513–37. pmid:21801010
  38. 38. Gonzalez-Reyes LE, Verbitsky M, Blesa J, Jackson-Lewis V, Paredes D, Tillack K, et al. Sonic hedgehog maintains cellular and neurochemical homeostasis in the adult nigrostriatal circuit. Neuron. 2012;75(2):306–19.PMC3408586. pmid:22841315
  39. 39. Machold R, Hayashi S, Rutlin M, Muzumdar MD, Nery S, Corbin JG, et al. Sonic hedgehog is required for progenitor cell maintenance in telencephalic stem cell niches. Neuron. 2003;39(6):937–50. pmid:12971894
  40. 40. Goodrich LV, Milenkovic L, Higgins KM, Scott MP. Altered neural cell fates and medulloblastoma in mouse patched mutants. Science. 1997;277(5329):1109–13. pmid:9262482
  41. 41. Cole F, Krauss RS. Microform holoprosencephaly in mice that lack the Ig superfamily member Cdon. Curr Biol. 2003;13(5):411–5. pmid:12620190
  42. 42. Chuang PT, Kawcak T, McMahon AP. Feedback control of mammalian Hedgehog signaling by the Hedgehog-binding protein, Hip1, modulates Fgf signaling during branching morphogenesis of the lung. Genes Dev. 2003;17(3):342–7.PMC195990. pmid:12569124
  43. 43. Bai CB, Joyner AL. Gli1 can rescue the in vivo function of Gli2. Development. 2001;128(24):5161–72. pmid:11748151
  44. 44. Garcia AD, Petrova R, Eng L, Joyner AL. Sonic hedgehog regulates discrete populations of astrocytes in the adult mouse forebrain. J Neurosci. 2010;30(41):13597–608.PMC2966838. pmid:20943901
  45. 45. Castillo-Azofeifa D, Losacco JT, Salcedo E, Golden EJ, Finger TE, Barlow LA. Sonic hedgehog from both nerves and epithelium is a key trophic factor for taste bud maintenance. Development. 2017;144(17):3054–65.PMC5611957. pmid:28743797
  46. 46. Donnelly CR, Kumari A, Li L, Vesela I, Bradley RM, Mistretta CM, et al. Probing the multimodal fungiform papilla: complex peripheral nerve endings of chorda tympani taste and mechanosensitive fibers before and after Hedgehog pathway inhibition. Cell Tissue Res. 2021;387(2):225–47. pmid:34859291
  47. 47. Lu W-J, Mann RK, Nguyen A, Bi T, Silverstein M, Tang JY, et al. Neuronal delivery of Hedgehog directs spatial patterning of taste organ regeneration. Proc Natl Acad Sci U S A. 2018;115(2):E200–E9. pmid:29279401
  48. 48. Kumari A, Ermilov AN, Allen BL, Bradley RM, Dlugosz AA, Mistretta CM. Hedgehog Pathway Blockade with the Cancer Drug LDE225 Disrupts Taste Organs and Taste Sensation. J Neurophysiol. 2015;113(3):1034–40.PMC4312875. pmid:25392175
  49. 49. Qin Y, Sukumaran SK, Jyotaki M, Redding K, Jiang P, Margolskee RF. Gli3 is a Negative Regulator of Tas1r3-Expressing Taste Cells. PLoS Genet. 2018;14(2):e1007058.PMC5819828. pmid:29415007
  50. 50. Smith RL, Geller AI, Escudero KW, Wilcox CL. Long-term expression in sensory neurons in tissue culture from herpes simplex virus type 1 (HSV-1) promoters in an HSV-1-derived vector. J Virol. 1995;69(8):4593–9.PMC189257. pmid:7609023
  51. 51. Nakayama A, Miura H, Shindo Y, Kusakabe Y, Tomonari H, Harada S. Expression of the basal cell markers of taste buds in the anterior tongue and soft palate of the mouse embryo. J Comp Neurol. 2008;509(2):211–24. pmid:18465790
  52. 52. Kumari A, Mistretta CM. Anterior and Posterior Tongue Regions and Taste Papillae: Distinct Roles and Regulatory Mechanisms with an Emphasis on Hedgehog Signaling and Antagonism. Int J Mol Sci. 2023;24(5).PMC10002505. pmid:36902260
  53. 53. Miura H, Kusakabe Y, Harada S. Cell lineage and differentiation in taste buds. Arch Histol Cytol. 2006;69(4):209–25. pmid:17287576
  54. 54. Peterson SC, Eberl M, Vagnozzi AN, Belkadi A, Veniaminova NA, Verhaegen ME, et al. Basal cell carcinoma preferentially arises from stem cells within hair follicle and mechanosensory niches. Cell Stem Cell. 2015;16(4):400–12.PMC4387376. pmid:25842978
  55. 55. Xiao Y, Thoresen DT, Williams JS, Wang C, Perna J, Petrova R, et al. Neural Hedgehog signaling maintains stem cell renewal in the sensory touch dome epithelium. Proc Natl Acad Sci U S A. 2015;112(23):7195–200. pmid:26015562
  56. 56. Brownell I, Guevara E, Bai CB, Loomis CA, Joyner AL. Nerve-derived sonic hedgehog defines a niche for hair follicle stem cells capable of becoming epidermal stem cells. Cell Stem Cell. 2011;8(5):552–65.PMC3089905. pmid:21549329
  57. 57. Jensen E. Technical review: In situ hybridization. Anat Rec (Hoboken). 2014;297(8):1349–53. pmid:24810158
  58. 58. Iwasaki S, Okumura Y, Kumakura M. Ultrastructural study of the relationship between the morphogenesis of filiform papillae and the keratinization of the lingual epithelium in the mouse. Cells Tissues Organs. 1999;165(2):91–103. pmid:10516422
  59. 59. Cobourne MT, Miletich I, Sharpe PT. Restriction of sonic hedgehog signalling during early tooth development. Development. 2004;131(12):2875–85. pmid:15151988
  60. 60. Echevarría-Andino ML, Franks NE, Schrader HE, Hong M, Krauss RS, Allen BL. CDON contributes to Hedgehog-dependent patterning and growth of the developing limb. Dev Biol. 2023;493:1–11. pmid:36265686
  61. 61. Izzi L, Lévesque M, Morin S, Laniel D, Wilkes Brian C, Mille F, et al. Boc and Gas1 Each Form Distinct Shh Receptor Complexes with Ptch1 and Are Required for Shh-Mediated Cell Proliferation. Dev Cell. 2011;20(6):788–801. pmid:21664577
  62. 62. Lee CS, Fan C-M. Embryonic expression patterns of the mouse and chick Gas1 genes. Mech Dev. 2001;101(1–2):293–7. pmid:11231094
  63. 63. Tenzen T, Allen BL, Cole F, Kang JS, Krauss RS, McMahon AP. The cell surface membrane proteins Cdo and Boc are components and targets of the Hedgehog signaling pathway and feedback network in mice. Dev Cell. 2006;10(5):647–56. pmid:16647304
  64. 64. Lee CS, Buttitta L, Fan CM. Evidence that the WNT-inducible growth arrest-specific gene 1 encodes an antagonist of sonic hedgehog signaling in the somite. Proc Natl Acad Sci U S A. 2001;98(20):11347–52.PMC58732. pmid:11572986
  65. 65. Del Sal G, Ruaro ME, Philipson L, Schneider C. The growth arrest-specific gene, gas1, is involved in growth suppression. Cell. 1992;70(4):595–607. pmid:1505026
  66. 66. Seppala M, Thivichon-Prince B, Xavier GM, Shaffie N, Sangani I, Birjandi AA, et al. Gas1 Regulates Patterning of the Murine and Human Dentitions through Sonic Hedgehog. J Dent Res. 2022;101(4):473–82.PMC8935464. pmid:34796774
  67. 67. Lan Y, Jiang R. Sonic hedgehog signaling regulates reciprocal epithelial-mesenchymal interactions controlling palatal outgrowth. Development. 2009;136(8):1387–96.PMC2687468. pmid:19304890
  68. 68. Jeong J, Mao J, Tenzen T, Kottmann AH, McMahon AP. Hedgehog signaling in the neural crest cells regulates the patterning and growth of facial primordia. Genes Dev. 2004;18(8):937–51. pmid:15107405
  69. 69. Parada C, Chai Y. Mandible and tongue development. Cur Top Dev Biol. 2015;115:31–58. pmid:26589920
  70. 70. Yu W, Ishan M, Yao Y, Stice SL, Liu HX. SOX10-Cre-Labeled Cells Under the Tongue Epithelium Serve as Progenitors for Taste Bud Cells That Are Mainly Type III and Keratin 8-Low. Stem Cells Dev. 2020;29(10):638–47.PMC7232695. pmid:32098606
  71. 71. Xu J, Liu H, Lan Y, Jiang R. The transcription factors Foxf1 and Foxf2 integrate the SHH, HGF and TGFbeta signaling pathways to drive tongue organogenesis. Development. 2022;149(21).
  72. 72. Zhang W, Yu J, Fu G, Li J, Huang H, Liu J, et al. ISL1/SHH/CXCL12 signaling regulates myogenic cell migration during mouse tongue development. Development. 2022;149(21):dev200788. pmid:36196625
  73. 73. Agbulut O, Noirez P, Beaumont F, Butler-Browne G. Myosin heavy chain isoforms in postnatal muscle development of mice. Biol Cell. 2003;95(6):399–406. pmid:14519557
  74. 74. Jiang Y, Du Z, Chen L. Histological study of postnatal development of mouse tongues. Exp Ther Med. 2018;15(1):383–6. pmid:29375694
  75. 75. Audu G, Kumari A. Novel Expression and Roles of Hedgehog Co-Receptor Gas1 during Embryonic Tongue Development. The Association for Chemoreception Sciences; April 21;2023 Bonita Springs.
  76. 76. Ishikawa Y, Ida-Yonemochi H, Saito K, Nakatomi M, Ohshima H. The Sonic Hedgehog–Patched–Gli Signaling Pathway Maintains Dental Epithelial and Pulp Stem/Progenitor Cells and Regulates the Function of Odontoblasts. Front Dent Med. 2021;2.
  77. 77. Mistretta CM, Kumari A. Tongue and Taste Organ Biology and Function:Homeostasis Maintained by Hedgehog Signaling. Annu Rev Physiol. 2017;79 24.1–2.PMC5966821. pmid:28192057
  78. 78. Rice R, Connor E, Rice DP. Expression patterns of Hedgehog signalling pathway members during mouse palate development. Gene Expr Patterns. 2006;6(2):206–12. pmid:16168717
  79. 79. Okubo T, Clark C, Hogan BLM. Cell lineage mapping of taste bud cells and keratinocytes in the mouse tongue and soft palate. Stem Cells. 2009;27(2):442–50. pmid:19038788
  80. 80. Gritli-Linde A. Molecular control of secondary palate development. Dev Biol. 2007;301(2):309–26. pmid:16942766
  81. 81. Lan Y, Jiang R. Sonic hedgehog signaling regulates reciprocal epithelial-mesenchymal interactions controlling palatal outgrowth. Development. 2009;136(8):1387–96. PMC2687468. pmid:19304890
  82. 82. Rice R, Spencer-Dene B, Connor EC, Gritli-Linde A, McMahon AP, Dickson C, et al. Disruption of Fgf10/Fgfr2b-coordinated epithelial-mesenchymal interactions causes cleft palate. J Clin Invest. 2004;113(12):1692–700. pmid:15199404
  83. 83. Mistretta CM, Kumari A. Hedgehog Signaling Regulates Taste Organs and Oral Sensation: Distinctive Roles in the Epithelium, Stroma, and Innervation. Int J Mol Sci. 2019;20(6).PMC6471208. pmid:30884865