Abstract
The current study focused on the histogenesis of the esophagus in quail embryos. Formation of the gut tube occurred on the 4th day of incubation. Development of the muscular layers occurred in a sequential manner; the inner circular layer on the 7th day, the outer longitudinal layer on the 8th day and the muscularis mucosae on the 9th day. Glandular development began on the 13th day of incubation. The epithelium was pseudostratified columnar that consisted of mucous cells, dendritic cells, and keratinocyte precursors. Epithelial stratification occurred on the 15th day of incubation. We used Mallory trichrome, Weigert-Van Gieson, and Gomori silver stains to visualize fibrous components. Scanned samples showed formation of endoderm and mesoderm on the 5th day of incubation. A layer of myoblasts developed on the 8th day of incubation. Formation of mucosal folds, which contained glandular openings, occurred on the 14th to 17th days of incubation. On the 5th to 8th days of incubation, CD34 and vascular endothelial growth factor (VEGF) positive-mesodermal cells, and telocytes (TCs) were detected. On the 15th day of incubation, CD34 and VEGF positive-telocytes, and fibroblasts, were identified. The current study described the correlations between functional morphology and evolutionary biology.
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Introduction
The esophagus is a muscular tube that connects the pharynx to the stomach. The major differences between the avian and the mammalian esophagus are the presence of glandular lamina propria and absence of voluntary muscular movement in birds1,2,3. The avian esophagus consists of the mucosal, submucosal, and muscular layers, and adventitia. Differences in histological features of the esophagus among birds have been identified3,4.
The quail is considered as an ideal animal model for use in developmental biology5. They are easily bred in the laboratory, have a short incubation period, and rapidly reach maturation6. The genetic profile of the Japanese quail is almost similar to the chicken7. Quails have also been recently introduced as an effective laboratory bird to be used for the production of transgenic avian8.
Evolutionary developmental biology (EvoDevo) is defined as the acquired new morphological characters that allows the assumption of a new function responding to the environmental conditions “adaptive zone”9. Evo‐devo aims to realize the origin and evolution of embryonic development including the mechanism of development and developmental modulation that driven the morphological novelty, the adaptive response of development in life-history evolution, the effect of ecological changes on development to permit evolutionary modification and the developmental and phylogenetic regulations of homoplasy and homology10. In the current study, we focused on the morphological novelties of quail esophagus including absence of ciliated epithelium and striated muscles in quail esophagus, alternative development of surface mucous cells/esophageal glands, short glands and shallow submucosa. We also the compared the morphological features of quail esophagus to other avian, mammalian, fish, reptilian species and discussed the possible role in adaptation to their environments.
Determining the histogenesis of different organs is necessary to gain insight and basic knowledge about the accurate timing of the histological events that occur during embryonic development. Quails may be used to study esophageal motility disorders such as achalasia and esophageal atresia. The current study focused on the histological development of the esophagus in quail embryos.
Material and methods
Sample collection and preparation of paraffin sections
Fertile quail (Coturnix Coturnix japonica) eggs were obtained from a farm belonging to the Department of Histology, Faculty of Veterinary Medicine, South Valley University, Qena, Egypt. Incubation of eggs was performed according to protocol11. Eggs were incubated at 37.5 °C with a relative humidity 65%. Quail embryos were collected from the 4th to the 17th days (final day) of incubation. Embryonic stages were determined upon the onset of incubation.
Sample collection and fixation for preparation of paraffin embedding were performed according to instructions11. The eggs were opened from the wide end by using a hand scalpel, then the embryos were carefully pulled from the egg shells. We used 2 petri dishes; one during egg opening and the other for placing the embryos, which were washed with distilled water then saline solution (0.9% NaCl). Embryos at days 3–13 of incubation were kept at minus 20 °C for 4 h prior to collecting, while days 15–17 were sacrificed and their skin incised from the recti of beak to the thoracic inlet, allowing access to the body cavity. Sample collection was done following the guidelines of the Institutional Ethical Committee of Veterinary Medicine, South Valley University, Egypt, and following the Egyptian Animals' laws. Bouin’s fixative (Table 1)12 was used for embryo fixation. Five embryos were collected for each incubation age and immersed in Bouin’s solution for 24 h. Embryos at days 15 and 17 of incubation were decalcified using 10% ethylenediaminetetraacetic acid. The protocol of the study was approved by the Egyptian Research Ethics Committee. Whole embryos were processed for preparation of paraffin sections, and was performed according to instructions13. Samples were dehydrated using ascending grades of ethanol (Table 1), cleared using methyl benzoate (Table 1), and embedded in paraffin wax (Table 1). Paraffin sections were sliced using Richert Leica RM 2125 Microtome, Germany.
Histochemical staining
For histological examination, paraffin sections were stained by hematoxylin and eosin (H&E)14. Mallory trichrome stain was used to visualize collagen fibers and the muscles, and was performed according to instructions15. Bielschowsky’s silver stain was used to visualize myofilaments, and was performed following instructions as indicated16. Gomori silver stain was used for visualization of reticular fibers, and was performed according to instructions17. Weigert-Van Gieson stain was used according to instructions12.
The stained sections were examined using a Leitz Dialux 20 microscope. Photos were taken using a Canon digital camera (Canon Powershot A95).
Immunohistochemistry (IHC) staining using CD34
Samples used for IHC investigations were fixed in NBF (neutral buffered formalin) (Table 1). Detection of antigen localization was performed using a combination of the avidin–biotin complex technique18 and the reagent of the Ultra Vision Detection System (Anti-Polyvalent, HRP/DAB manufactured by Thermo Fisher Scientific TP-015HD). The procedures were performed according to the manufacturer’s instructions19,20,21. Paraffin sections measuring 5 µm were dewaxed by xylene, hydrated with ascending grades of alcohol, and washed for 5 min twice with PBS (phosphate buffered saline) at pH 7.4 (Table 1). The sections were treated with 3% hydrogen peroxide in methanol at room temperature for 20 min to block endogenous peroxidase activity. The sections were rinsed for 10 min with running tap water. Improving antigen retrieval requires use of a 10 mm sodium citrate buffer (pH 6.0) (Table 2) in a water bath for 20 min at 95–98 °C. This was followed by slide cooling for 20 min at room temperature, and subsequently washed for 5 min twice with PBS at pH 7.4. Ultra V block was used for 5 min at room temperature to prevent nonspecific background staining. To avoid artifacts, use of ultra V block should not exceed 10 min. The primary antibody (Table 3) was applied on the sections overnight at 4 °C. The sections were then washed for 5 min thrice with PBS at pH 7.4. The biotinylated secondary antibody (Goat Anti-Polyvalent, Anti-Mouse IgG + Anti -Rabbit IgG; Thermo Fisher Scientific, UK; Lab Vision Corporation; Table 2) was applied on the sections for 10 min at room temperature. The sections were then washed for 5 min twice with PBS at pH 7.4 and incubated for 10 min at room temperature using streptavidin-peroxidase complex (Thermo Fisher Scientific, UK; Lab Vision Corporation, USA). Visualization of the bound antibodies was carried out by incubating the section in a humid chamber using a mixture of 1 drop of 3,3′-diaminobenzidine (DAB) and chromogen (Table 4) to 2 mL of DAB plus substrate for 5 min at room temperature. The sections were counterstained with Harris hematoxylin for 30 s. The sections were dehydrated using ethanol and isopropanol I and II, cleared in xylene, and covered by DPX. CD34 immunohistochemical staining was examined using the Leitz Dialux 20 microscope provided with the Canon (Power shot A95) digital camera.
Immunohistochemical procedures of vascular endothelial growth factor (VEGF)
Samples used for VEGF immunostaining were fixed in NBF (neutral buffer formalin) (Table 1). The two-step immunohistochemical staining procedures utilized the DAKO EN Vision TM and System, HRP peroxidase22. The staining was performed according to instructions23. Paraffin sections of 5 µm were dewaxed, rehydrated, and rinsed for 5 min twice with PBS at pH 7.4. Blocking of the endogenous peroxidase activity was achieved by using drops of 3% hydrogen peroxide in methanol for 20 min at room temperature, then thoroughly washed with running tap water for 10 min. Antigen retrieval was performed using 10 mm sodium citrate buffer (pH 6.0) (Table 2). The buffer was heated to 95 °C–98 °C in a water bath for 20 min, followed by cooling at room temperature for 20 min. Sections were washed for 5 min twice with PBS at pH 7.4. Blocking nonspecific background staining was performed using drops of blocking serum (DAKO) to cover the sections for 5 min at room temperature. The sections were then incubated with the primary antibody. The antibodies that were applied showed immunoreactivity in avian species24. Table 3 explored the identity, sources, and the working dilution of antibodies used in the immunohistochemical technique. The slides were rinsed for 5 min thrice with PBS at pH 7.4 then incubated with secondary antibody for 30 min at room temperature. The slides were again washed for 5 min twice with PBS at pH 7.4 and incubated for 5–10 min at room temperature with 3,3′-diaminobenzidine (DAB) and substrate-chromogen that produced a brown color at the antigen site. The slides were then counterstained with Harris hematoxylin for 30 s. The sections were dehydrated with ethanol alcohol 90%, then 100% II, cleared in xylene, and covered using DPX. VEGF immunohistochemically-stained sections were examined using the Leitz Dialux20 microscope provided with the Canon (PowerShot A95) digital camera -vecontrols were performed with the same procedures except using primary antibodies.
Preparation of samples for embedding in resin
Resin embedding technique was performed according to25 using Karnovsky’s fixed samples. Five samples were used from incubation days 5, 8, and 15. Each esophagus, measuring 2–3 mm in length, was carefully excised after sacrificing the quail. Karnovsky fixative26 was prepared from a mixture of 10 mL of 25% paraformaldehyde, 10 mL of 50% glutaraldehyde, 50 mL phosphate buffer, and 30 mL distilled water. Karnovsky fixative was applied overnight at 4 °C. The samples were post-fixated by osmium tetroxide, dehydrated using ascending grades of alcohol, inoculated in a mixture of alcohol/resin and pure resin, resin embedding and crystallization was performed in oven at 60 °C degrees. Semi-thin sections (1 μm) were created using an ultramicrotome (Ultracut E, Reichert-Leica, Germany) and stained with toluidine blue27,28, methylene blue29,30, and periodic acid-Schiff (PAS)31. Staining of semi-thin sections required dissolving of resin using saturated alcoholic solution of sodium hydroxide. The stained sections were examined using a Leitz Dialux 20 microscope and a Canon digital camera (Canon PowerShot A95).
Transmission electron microscopy
Ultra-thin Sects. (60 nm) of the esophagus at incubation days 8, 13, and 15 were taken by using a Reichert ultramicrotome. The sections were stained with uranyl acetate for 15 min then lead citrate for another 15 min. The stained grids were examined by using a JEOL100CX II transmission electron microscope at the central laboratory of South Valley University, Egypt.
Scanning electron microscopy
The samples were washed several times with normal saline and then fixed in 4% glutaraldehyde solution for 24 h at pH 7.3. They were then washed in sodium phosphate buffer at pH 7.3 4 times for 15 min and fixed in 1% osmic acid and 0.1 M sodium phosphate buffer for 2 h at room temperature. Thereafter, they were washed with 0.1 M sodium phosphate buffer 4 times for 15 min. The samples were then dehydrated with 50%, 70%, and 90% alcohol for 30 min in each concentration, then 100% for 2 days followed by isoamyl acetate for another 2 days. The samples were subjected to the critical point drying method (Critical Point Drying Procedure Polaron E3000 CPD Apparatus, Germany) and coated with gold using JEOL-1100 E-ion Sputtering Device (Japan). Samples were examined using a JEOL scanning electron microscope (JSM 5500 LV) at 10 kV at the central laboratory of South Valley University, Egypt.
Coloring of TEM and SEM images
We colored the images produced from the transmission (TEM) and scanning (SEM) electron microscopes using the Photo Filter 6.3.2 program to recognize different types of cells and to distinguish different structures. The methods used were as previously described19,30,32,33,34,35,36,37,38,39,40,41,42,43,44,45,46,47,48,49,50.
Ethical approval
The National Ethics Committee of South Valley University and veterinary authorities in South Valley University Province, Egypt, approved the method of this study. ‘All procedures were performed in accordance with the relevant guidelines and regulations’51. We stated that the study was carried out in compliance with the ARRIVE guidelines in Ethics approval section.
Results
On the 4th day of incubation, the gut tube was formed and consisted of endoderm and covered by mesoderm (Fig. 1A,B). On the 5th day of incubation, the primitive esophagus had an endodermal layer of pseudostratified epithelium and surrounded by a condensed layer of mesenchyme, a marker for the development of a muscular wall (Fig. 1C,D). This muscular wall began to develop on the 6th day of incubation. A circular layer of SMC (smooth muscle cells) was observed (Fig. 1E,F). On the 7th day of incubation, the circular muscular layer became more distinct while myoblasts aggregated to form the primitive outer longitudinal muscular layer (Fig. 2A–C). On the 8th and 9th days of incubation, the esophagus developed distinct inner circular and outer longitudinal muscular layers (Figs. 2A–I). The muscularis mucosae recognized on the 8th day of incubation by mesenchymal condensation and identified as a thin layer of SMC that supports the mucosa on the 9th day of incubation. (Fig. 2H). On the 13th day of incubation, the esophagus developed muscualris mucosae rather than the distinctive inner circular and outer longitudinal muscular layers (Fig. 3A–D). The epithelial invaginations formed sac-like glandular units. The epithelium was pseudostratified type (Fig. 3A,B). Presence of collagen fibers in the lamina propria, between the muscle fibers, and the serosa were visualized using Mallory trichrome (Fig. 3C) and Weigert-Van Gieson stains (Fig. 3D,E).
On the 15th day of incubation, reticular fibers were recognized using Gomori’s silver stain in the lamina propria (Fig. 4A,B), around the gland (Fig. 4D), around smooth muscle cells and in the myenteric or Auerbach's plexus (Fig. 4E,F). TCs were identified in the LP (Fig. 4C) and between the muscle bundles (Fig. 4G–I). Bielschowsky's silver stain was used to visualize the SMC, which revealed a granular appearance indicating the presence of dense bodies (Fig. 4G–I). The thoracic portion of the esophagus developed highly folded mucosa which exhibited numerous sac-like glandular units, inactive glands (Fig. 5A,B) the epithelial lining had stratified squamous epithelium non-keratinized (Fig. 5C). The cervical portion also had stratified squamous epithelium non-keratinized developed active mucous esophageal glands (Fig. 5D–F). Telocytes were recognized in the lamina propria (Fig. 5E) and submucosa (Fig. 5F).
The pseudostratified epithelium and the basal lamina appreaed PAS-positive on the 8th day of incubation (Fig. 6A–C). The developing muscular layers were distinguished by the myoblasts (Fig. 6D), some of which contained PAS-positive glycogen granules. PAS-positive myelin sheaths were also detected between myoblasts (Fig. 6D). On the 15th day of incubation, the thoracic portion of the esophagus was lined by pseudostratified epithelium exhibited PAS activity while the gland was inactive PAS-ve (Fig. 6E–G,J). PAS-ve collagen fibers were identified in the submucosa and muscular layer (Fig. 6G,K). Telocytes exhibited high affinity for PAS (Fig. 6G). The esophageal mucous glands in the cervical portion were now active and contained PAS-positive granules (Fig. 6H,I,L,M). On the 13th day of incubation, epithelial invaginations established sac-like glandular units. The epithelium was pseudostratified type (Fig. 7A–C) and had mucous-secreting inclusions that appreaed metachromatic by toluidine blue but the gland was inactive exhibited -ve PAS reaction (Fig. 7A–E). Mitotic figures were detected in the interstitial cells (Fig. 7B), epithelial cells (Fig. 7C), and SMC (Fig. 7F). On the 15th day of incubation, the esophagus exhibited distinct mucosal foldings, and the epithelium was now stratified squamous non-keratinized. (Fig. 7G,H). The esophageal mucous glands in the cervical portion were now active (Fig. 7G,H). TEM imaging revealed that the pseudostratified epithelium had superficial mucous-secreting cells (Fig. 8A–D) and the myoblasts developed (Fig. 8B). Most epithelial cells contained RER, ribosomes, and mitochondria (Fig. 8E), and putative dendritic cells had vesicles and lamellar granules (Fig. 8F). The lamina propria contained interstitial cells rich in dilated RER (Fig. 8G). The developing muscular layers were distinguished by the myoblasts (Fig. 8H). On the 17th day of incubation, the esophageal keratinocytes developed keratin intermediate filaments (Fig. 9A,B). Dendritic cells appeared between keratinocytes. Dendritic cells were recognized by vesicles, multivesicular bodies, dense granules, and rod-shaped granules (Fig. 9C). Esophageal glands contained mucous granules, RER, and exhibited interdigitation (Fig. 9D,E). Collagen fibers were identified using TEM (Fig. 9A,B,F).
On the 5th day of incubation, scanned samples of the esophagus showed the presence of endoderm and mesoderm and the endoderm was pseudostratified epithelium (Fig. 10A,B). On the 8th day of incubation, the pseudostratified epithelium was also existed (Fig. 10C–F). Development of the myoblast layer occurred at this age (Fig. 10E–H). On the 14th day of incubation, the esophagus formed a distinct mucosal fold which contained the opening of the esophageal glands (Fig. 10I–L), and on the 17th day of incubation (Fig. 10Q,S,T) the esophageal glands were now visible in the lamina propria (Fig. 10R). TCs were identified in the muscular layer (Fig. 10 M–P).
On the 5th day of incubation, CD34 and VEGF positive cells were identified in the subepithelial tissue and the mesenchyme. CD34 positive TCs were distinguished by cell prolongations or telopodes and VEGF positive TCs were recognized as well (Fig. 11A,B). On the 8th day of incubation, it is notable that CD34 positive cells were more obvious in the lamina propria compared to VEGF. CD34 and VEGF positive cells were also identified in the peri-muscular tissue. CD 34 positive and VEGF positive TCs were distinguished by their telopodes (Fig. 12A–C). The sprouting endothelial cells were CD34 positive (Fig. 13E). Figure 13 F represented negative control for CD34.
On the 15th day of incubation, Interstitial cells in the lamina propria and the submucosa were CD34 positive including TCs that had distinct telopodes, and fibroblasts which had voluminous cytoplasm and short cell processes (Fig. 13A–D). Perivascular tissue between muscles had strong CD34 immunoaffinity (Fig. 13D). CD34-positive TCs were detected between muscle fibers and bundles (Fig. 13D). Interstitial fibroblast-like cells in the lamina propria were VEGF positive (Fig. 14A). Endothelial sprouting in the lamina propria were VEGF positive (Fig. 14B). VEGF-positive TCs were identified in the lamina propria and the submucosa (Fig. 14C,D). Perivascular tissue had strong immunoaffinity for VEGF (Fig. 14E). Figure 14 F represented negative control for VEGF.
All events that occurred during esophageal development were summarized in (Fig. 15).
Discussion
The current study investigated histological events occurring during the development of the esophagus of the quail.
The gut tube was formed on the 4th day of incubation. This was composed of endoderm and covered by mesoderm, which was previously described by another study52. The authors mentioned the vacuolated stage, in which extracellular vacuoles appeared between the epithelial cells, and they suggested that the vacuoles may be related to the developmental cysts.
Mesenchymal condensation was the first sign of development of the muscular layer. Development of the muscular layer occurred in a sequential manner; the inner circular layer on the 7th day, the outer longitudinal layer on the 8th day, and the muscularis mucosae began by mesenchymal condensation and development of strands of SMC on the 9th day. A similar sequence of muscular development occurs in the pheasant esophagus; development of the inner circular layer of the begins on the 8th day, the outer longitudinal layer on the 12th day, and the muscularis mucosae is completed on the 18th day53. The sequence of muscular coat development of the human embryos is also similar to avian species. However, in the Guinea Fowl (Numida meleagris). The tunics of the muscularis are not well developed till at posthatch54. The muscular layers of the human esophagus develop in similar sequence to quail. In the 12 mm and 20 mm crown-rump stages human embryos, the inner muscular layer become more developed. The outer longitudinal muscle layer and the muscularis mucosae are prominent55.
In the current study, PAS-positive inclusions were detected in some myoblasts and myelin sheaths. PAS is used to identify the simple sugars in myelin56. The study used Bielschowsky's silver stain to visualize the SMC, which exhibited a granular appearance indicating dense bodies. Visualization of the myofilaments striations in skeletal muscles was documented by using Bielschowsky’s stain57.
Esophageal epithelium directly transformed from pseudostratified columnar into the stratified squamous non-keratinized type by the 15th day. In all stages of esophageal development, the epithelium was non-ciliated that may be a unique feature of quail esophagus. A different types of epithelium is noted in the pheasant; pseudostratified columnar epithelium is observed on the 9th day of incubation and transformed into stratified cuboidal by the 10th day, then to simple cuboidal epithelium between the 10th and 12th days, and finally to stratified squamous on the 13th day53. In the chukar partridge, the epithelium transformed to 2 layered cuboidal cells with some cilia by day 9 of incubation. The ciliated cells increased during embryonic development on day 10, 11. By day 18 of incubation, two types of simple and pseudostratified columnar epithelial tissue are detectable in the thoracic portion of esophagus. The cilia were difficult to distinguished due to the epithelial secretion58. Developmental variations of the esophagus are documented in human embryos. The esophageal epithelium is stratified columnar of four cells deep in the 8.5 mm crown-rump human embryo and become multilayered columnar in the 40 mm crown-rump stage. Epithelium is surrounded by undifferentiated mesenchyme and condensed cells form the inner muscle coat in the 8.5 mm crown-rump embryo. In the 30 to 40 mm crownrump embryo, the basal epithelial cells protrude toward the lumen to become ciliated columnar cells. The ciliated cells develop in the middle third of the esophagus and extend towards the cranial and caudal portions. The ciliated cells distribute along the entire mucosa of the esophagus of the 60 mm crown-rump embryo, except for the upper and lower ends. The ciliated cells gradually disappear with proceeding the development and only isolated patches of ciliated columnar cells occasionally remain until after birth55. Development of ciliated esophageal epithelium is associated with transformation into stratified columnar in type. Formation of ciliated stratified columnar epithelium occurs on the 8th week of gestation and were decreased after the14th week. The stratified columnar epithelium transforms into the stratified squamous epithelium during the 4th month59.
Glandular development began on the 13th day of incubation. Epithelium invaginated into the underlying connective tissue, forming sac-like glandular units. A similar occurrence can be observed in chickens, starting on the 12th day of incubation60. In pheasants, glandular development begins as epithelial buds in the esophagus on the 18th day of incubation53. The current study revealed epithelial transformation preceded glandular activation. The undifferentiated pseudostratified epithelial cells the differentiated into keratinocytes and dendritic cells. Esophageal glands which synthesize polysaccharide components were detected using PAS and toluidine blue. Through the metachromatic reaction from toluidine blue, some of these components were identified as glycosaminoglycans (GAGs), which the avian esophageal glands secrete along with mucopolysaccharides53,60. The submucosa of the quail esophagus was a delicate layer. Similar to other avian species, very thin layer of submucosa is described in the pariah kite, median egret, goshawk, dove and duck61, the House Sparrow62, and chukar partridge (Alectoris chukar)58. In pigeons, the submucosa is thin in female (40.2±7.5 in the cervical region and 20.6± 3.6 in the thoracic region) and is relatively thicker in male (60.7±11.5 in the cervical region and 40.6±8.9 in the thoracic region)63. While, the submucosa in the esophagus of human64 and small animals65 is a thick layer.
In the current study, we focused on the morphological novelties of quail esophagus and discussed the possible role in adaptation to their environments. Embryonic esophageal epithelium of quail didn’t acquire cilia. Unlike, other avian and mammalian species55,58,59. The development of the cilia is related to the functional requirements. It is not clear whether the cilia are motile true cilia that remove the surface secretions/foreign particles or non-motile stereocilia that has absorptive function. In chick embryos, the esophageal cilia associate with microvilli and microplicae66. Esophageal cilia have been described as transitory structures in avian and mammals in contrast to the lower vertebrates such as the crocodiles. Esophagus of the juvenile American alligators (Alligator mississippiensis) has ciliated columnar epithelium67. It is interesting that embryonic quail esophagus lacked the skeletal muscles, unlike the higher (mammalian) and lower species (aquatic and reptiles). The esophagus is devoid of striated muscles in wild avian species such as Rock Dove, Collared Dove, Rose-ringed Parakeet, Kestrel, House Sparrow and Linnet68, mallard, spot-billed duck, Ural owl and Hodgson's hawk-eagle69, and other birds, Kingfisher (Halcyon smyrnensis) and Hoopoe (Upupa epops)70 and chicken71. Thus, it seems that involuntary control of the esophagus is unique feature of avian species. This may be correlated to feeding habitat that requires peristatic involuntary movement during regurgitative feeding to feed their young. It seems that regurgitation may require definite time lapsing to deliver a semi-digested food. In mammals, the muscular wall of the esophagus is comprised of both skeletal and smooth muscles. However the distribution of the two types varies according to animal species72,73. Among fish, the esophagus of Anablepsoidesurophthalmus has some striated muscle fibers74. While, heavy striated musculature thick inner longitudinal and outer circular layer are described in tilapia75, Trachelyopterus striatulus (Siluriformes: Auchenipteridae)76. Krauss et al. explain the development of the esophageal muscular layer with the variable degree of esophageal striated myogenesis. Esophageal striated muscle progenitor cells that originate in the craniopharyngeal mesoderm and express the early marker Mesp1, and subsequently express Tbx1 and Isl1. Esophageal striated muscle progenitor cells colonize the proximal portion of the esophageal muscular layer. They migrate distally along the esophageal muscular layer forming the transition zone where they express Pax7, Myf5 and MyoD, and myogenin. Some Pax7+ and Myf5+/MyoD+ cells propagate to generate adequate numbers of precursor cells for the muscular layer. Some differentiate into striated myofibers, which form proximal to the transtion zone. The authors descried a linear expression pattering Isl1, Pax7, Myf5/MyoD progression77.
The architecture of the esophageal glands of avian and mammals are quite similar. Mammalian esophageal glands are compound tubuloalveolar type in mammals73 and some birds such as barn owl (Tyto alba) and common wood pigeon78, and compound tubular type in wild bronze turkey (Meleagris gallopavo)79. The avian esophageal glands modify that have short ducts, and located in the lamina propria. While, mammalian esophageal glands are more developed dominated the submucosa69. The ducting system have some of characteristics different according animals species73. It is noted that esophageal mucous glands progressively develop with higher classes of the evolution tree. The geometric parameters of esophageal submucosal glands are measured in the avian and porcine species. The esophageal submucosal glands comprised 35% in avian and 45% in porcine area of the submucosa. The glands have an area of 125,000 μm2 in avian) and 580,000 μm2 in porcine80. Some fish esophagi lack the esophageal glands, instead mucous cells are located in the epithelium. This occurs in Sphoeroides testudineus81, Trichomycterus brasiliensis82 and Anablepsoidesurophthalmus74, Nile tilapia and African catfish75, Larimichthys crocea (Acanthopterygii: Perciformes)83. While, other aquatic fish, the Asian seabass (Lates calcarifer), has both goblet cells and esophageal glands84. In crocodiles, the juvenile American alligators (Alligator mississippiensis), the esophageal epithelium has few mucous goblet cells67. The mucous secretion is essential to moisten food, lubrication and have protective function to neutralize the gastric acidity in case gastric reflux85. The evaluation of the stomach and the specification of gastric epithelium is described among fishes, amphibians, reptiles, birds and mammals86. Comparisons of stomach acidity in mammal and bird indicates that scavengers and carnivores exhibit higher stomach acidities compared to herbivores or carnivores feeding on phylogenetically distant prey87. Delicate submucosa is a characteristic feature of avian esophagus. This may be related to absence of the submucosal glands. unlike mammals, the abundant glandular subunits occupied the submucosa in dogs while extends to the tunica muscularis in rabbit73.
The current study detected localization of collagen and reticular and elastic fibers in embryonic quail esophagus. Staining with Mallory trichrome and Weigert-Van Gieson revealed the presence of collagen fibers in the basal lamina, lamina propria, between the muscle fibers, and the serosa. Reticular fibers were detected in the lamina propria, myocytes, around glands, and in the myenteric or Auerbach's plexus. The observation of reticular fibers around the SMC was supported by previous studies88. Van-Geison stain was used to detect collagen in the pheasant53 and geese89 esophagus, particularly in the lamina propria and submucosa of the latter. The amount of collagen fibers in the avian esophagus depended on the feeding habits of the species. The lamina propria had more collagen fibers in pariah kites, egrets, and doves, while only a few collagen and reticular fibers were detected in goshawks and ducks. Collagen and elastic fibers are found in the tunica adventitia of the esophagus of pariah kites, median egrets, goshawks, doves, and ducks61.
Putative dendritic cells were found in the pseudostratified epithelium. Vesicles and granules were observed on the 8th day of incubation. Active dendritic cells were identified after epithelial stratification, and they contained vesicles, multivesicular bodies, dense granules, and rod-shaped granules. Dendritic cells act as antigen-presenting cells and contribute to the adaptive and innate immune response76,77,92. Dendritic cells were recognized using TEM by the presence of the dendritic lamellar and multivesicular bodies93,94. They also express MHC class II antigen which stimulates naïve T-lymphocytes and initiates the primary immune response. They are also involved in the activation of B-cells and reactivation of the memory B lymphocytes90,92.
In the current study, we used CD34, a common marker for undifferentiated stem cells, to demonstrate the distribution of different types of putative stem cells during the different stages of esophageal development. On the 5th day of incubation, CD34 positive cells were identified in the, subepithelial tissue, and mesenchyme, while on the 8th day they were also identified in the lamina propria and peri-muscular tissue. On the 15th day. The, interstitial cells including TCs and fibroblasts, exhibited CD34 immunoreactivity. Telocytes have an essential role in morphogenesis, embryogenesis, organization of the embryonic tissues81,82,97, cell migration, cell adhesion, proliferation, differentiation33,35,96, tissue homoeostasis, remodeling98 and repair99. The CD34 protein is a member of a family of transmembrane sialomucin proteins. CD34 is common for marker of hematopoietic stem cells, hematopoietic progenitor cells, as well as non-hematopoietic cells such as vascular endothelial progenitors and embryonic fibroblasts, multipotent mesenchymal stromal cells, interstitial dendritic cells, and epithelial progenitors. CD34 is critical for adhesion molecules100, proliferation and blocking differentiation of stem or progenitor cells87,88,103.
Angiogenesis is the physiological process of forming new blood vessels from the pre-existing vessels. Growth of the vasculature is associated with proliferation of the endothelial cells to form sprouting vascular branches. Angiogenic events are essential during development to provide adequate nutrients for the microenvironment required by cells and tissues. If the oxygen levels are low, cells may experience hypoxia. Hypoxic cells express vascular proliferation markers to induce and enhance angiogenesis104,105. In the current study, we used immunohistochemistry to demonstrate the types of cells that express VEGF, a vascular proliferation marker. On the 5th day of incubation, VEGF positive cells were identified in the subepithelial tissue, and mesenchyme, and on the 8th day of incubation, they were also identified in the lamina propria and peri-muscular tissue. VEGF positive TCs were distinguished by their telopodes. On the 15th day of incubation, Telocytes may promote endothelial cell proliferation and angiogenesis106. Stromal fibroblasts express VEGF107. VEGF is secreted by the salivary gland and also shed in the saliva. Salivary VEGF is an essential stimulus for oral mucosal tissue repair108,109.
Conclusion
The current study provided information on esophageal development, including a timeline of the development events, and described changes of cellular components. The quail esophagus may be used as a model to study esophageal disorders. The current study also discussed the unique morphological features of quail esophagus and analyze the evolutionary morphology among different classes in relation to the function.
Abbreviations
- SMC:
-
Smooth muscle cells
- NBF:
-
Neutral buffered formalin
- PAS:
-
Periodic acid-schiff stain
- TC:
-
Telocyte
- VEGF:
-
Vascular endothelial growth factor
References
Bacha, W. J. & Bacha, L. M. Color Atlas of Veterinary Histology 3rd edn. (Wiley-Blackwell, 2012).
Sturkie, P. D. Avian Physiology (Springer, 2012).
Reece, W. O., Erickson, H. H., Goff, J. P. & Uemura, E. E. Dukes’ Physiology of Domestic Animals (Wiley, Hoboken, 2015).
Rossi, J. R. et al. Morphology of oesophagus and crop of the partridge Rhynchotus rufescens (tiramidae). Acta Sci. Biol. Sci. 28, 165–168 (2006).
Ainsworth, S. J., Stanley, R. L. & Evans, D. J. Developmental stages of the Japanese quail. J. Anat. 216, 3–15. https://doi.org/10.1111/j.1469-7580.2009.01173.x (2010).
Huss, D., Poynter, G. & Lansford, R. Japanese quail (Coturnix japonica) as a laboratory animal model. Lab. Anim. 37, 513–519 (2008).
Hoeben, A. et al. Vascular endothelial growth factor and angiogenesis. Pharmacol. Rev. 56, 549–580. https://doi.org/10.1124/pr.56.4.3 (2004).
Poynter, G., Huss, D. & Lansford, R. Japanese quail: An efficient animal model for the production of transgenic avians. Cold Spring Harbor Protoc. https://doi.org/10.1101/pdb.emo112 (2009).
Bertossa, R. C. Morphology and behaviour: Functional links in development and evolution. Philos. Trans. R. Soc. Lond. B 366, 2056–2068. https://doi.org/10.1098/rstb.2011.0035 (2011).
Hall, B. Evo-devo or devo-evo: Does it matter. Evol. Dev. 2, 177–178 (2000).
Soliman, A. S. Potential role of telocytes in differentiation of embryonic skeletal progenitor cells. SF J Stem Cell. 1, 1 (2017).
Suvarna, K., Layton, C. & Bancroft, J. Bancroft’s Theory and Practice of Histological Techniques (Elsevier, 2013).
Soliman, S. A. Organization and pattering of mesenchymal cells in quail embryonic cartilage. SF J Stem Cell 1, 1 (2018).
Harris, H. On the rapid conversion of haematoxylin into haematein in staining reactions. J. Appl. Microsc. Lab. Methods 3, 777 (1900).
Mallory, F. B. The anilin blue collagen stain. Stain Technol. 11, 101 (1936).
Intorcia, A. et al. A Modification of the Bielschowsky Silver Stain for Alzheimer Neuritic Plaques: Suppression of Artifactual Staining by Pretreatment with Oxidizing Agents. (2019).
Suvarna, K. S., Layton, C. & Bancrof, J. D. Bancroft’s Theory and Practice of Histological Techniques: Expert Consult: Online and Print (Elsevier Health Sciences, 2012).
Hsu, S. M., Raine, L. & Fanger, H. Use of avidin-biotin-peroxidase complex (ABC) in immunoperoxidase techniques: A comparison between ABC and unlabeled antibody (PAP) procedures. J. Histochem. Cytochem. 29, 577–580 (1981).
Abd-Elhafeez, H. H., Abou-Elhamd, A. S. & Soliman, S. A. Morphological and immunohistochemical phenotype of TCs in the intestinal bulb of Grass carp and their potential role in intestinal immunity. Sci. Rep. 10, 14039 (2020).
Mustafa, F.E.-Z. & Elhanbaly, R. Distribution of estrogen receptor in the rabbit cervix during pregnancy with special reference to stromal elements: An immunohistochemical study 10. Sci. Rep. 10, 13655 (2020).
Abd-Elkareem, M. & Abou-Elhamd, A. S. Immunohistochemical localization of progesterone receptors alpha (PRA) in ovary of the pseudopregnant rabbit. Anim. Reprod. 16, 302–310 (2019).
Kämmerer, U. K. M. et al. A new rapid immunohistochemical staining technique using the EnVision antibody complex. J. Histochem. Cytochem. 49, 623–630 (2001).
Abdo, W., Ghattas, S., Sakai, H., Hagiwara, H. & Yanai, T. Assessment of proliferative activity by proliferative cell nuclear antigen (PCNA) and anti-bromodeoxyuridine (BrdU) immunolabeling in the tissues of Japanese Eels (Anguilla japonica). Turk. J. Fish. Aquat. Sci. 14, 413–419 (2014).
Chen, Q. et al. Sexual differences in cell proliferation in the ventricular zone, cell migration and differentiation in the HVC of juvenile Bengalese finch. PLoS ONE 9, e97403 (2014).
Spurr, A. R. A low-viscosity epoxy resin embedding medium for electron microscopy. J. Ultrastruct. Res. 26, 31–43 (1969).
Karnovsky, M. J. A formaldehyde-glutaraldehyde fixative of high osmolarity for use in electron microscopy. J. Cell Biol. 27, 137A-138A (1965).
Soliman, S. A., Ahmed, Y. A. & Abdelsabour-Khalaf, M. Histogenesis of the stomach of the pre-hatching quail: A light microscopic study. Anat. Sci. Int. 91, 407–418 (2016).
Soliman, S. & Emeish, W. Morphological alternations of intraepithelial and stromal telocytes in response to salinity challenges. bioRxiv https://doi.org/10.1101/115881 (2017).
Abd-Elhafeez, H. H. & Soliman, S. A. New description of telocyte sheaths in the bovine uterine tube: An immunohistochemical and scanning microscopic study. Cells Tissues Organs https://doi.org/10.1159/000452243 (2017).
Abdel-Maksoud, F. M., Abd-Elhafeez, H. H. & Soliman, S. A. Morphological changes of telocytes in camel efferent ductules in response to seasonal variations during the reproductive cycle. Sci. Rep. 9, 4507. https://doi.org/10.1038/s41598-019-41143-y (2019).
Soliman, S. A. Morphological and histochemical description of quail feather development. Anat. Record https://doi.org/10.1002/ar.24276 (2019).
Fatma El-Zahraa, A. M. & Abd-Elhafez, E. A. A histological, histochemical and ultrastructural characterization of uterine vessels at early stages of pregnancy. J. Histol. Histopathol. Res. 2, 41–47 (2018).
Soliman, S. Potential role of telocytes in differentiation of embryonic skeletal progenitor cells. SF J Stem Cell 1, 1 (2017).
Soliman, S. A. & Madkour, F. A comparative analysis of the organization of the sensory units in the beak of duck and quail. Histol. Cytol. Embryol https://doi.org/10.15761/HCE.1000122 (2017).
Soliman, S. A. Telocytes during organogenesis: Relations to nephrogenic cords in mesonephros of quail embryos. Histol. Cytol. Embryol. 1, 1–6 (2017).
Abd-Elhafeez, H. H., Abou-Elhamd, A. S., Abdo, W. & Soliman, S. A. Migratory activities and stemness properties of rodlet cells. Microsc. Microanal 21, 1–18. https://doi.org/10.1017/S1431927620001828 (2020).
Abd-Elhafeez, H. H., Hassan, A. H. S., & Hussein, M. T. Melatonin administration provokes the activity of dendritic reticular cells in the seminal vesicle of Soay ram during the non-breeding season. Sci. Rep. 11(1), 872 (2021).
Abd-Elhafeez, H. H., Abdo, W., Kamal, B. M., & Soliman, S. A. Fish telocytes and their relation to rodlet cells in ruby-red-fin shark (rainbow shark) Epalzeorhynchos frenatum (Teleostei: Cyprinidae). Sci. Rep. 10(1) (2020).
Anwar, S., Abd-Elhafeez, H.., Abdel-Maksoud, F. & Abdalla, K. Morph-anatomic and histochemical study of ileum of goose (Alopochen egyptiacus) with special references to immune cells, mucous and serous goblet cells, telocytes, and dark and light smooth muscle fibers. Microsc. Res. Tech. https://doi.org/10.1002/jemt.23692 (2021).
Soliman, S. A. Telocytes are major constituents of the angiogenic apparatus. Sci. Rep. 11(1), 5775 (2021).
Soliman, S. A., Kamal, B. M.,& Abd-Elhafeez H. H. Cellular invasion and matrix degradation, a different type of matrix-degrading cells in the cartilage of catfish (Clarias gariepinus) and Japanese quail embryos (Coturnix coturnix japonica). Microsc. Microanal. 25(05), 1283–1292 (2019).
Soliman, S. A., Kamal, B. M., Abuo-Elhmad, A. S., & Abd-Elhafeez, H. H. Morphological and histochemical characterization of the dermal plates of pleco (Hypostomus plecostomus). Microsc. Microanal 26, 551–566 (2020).
Abd-Elhafeez, H. H., & Soliman, S. A. Origin of rodlet cells and mapping their distribution in ruby-red-fin shark (Rainbow Shark) epalzeorhynchos frenatum (teleostei: cyprinidae): Light, immunohistochemistry and ultrastructure study. J. Cytol. Histol. 7, 435 (2016).
Soliman, S. A., & Abd-Elhafeez, H. H. Mesenchymal cells in cartilage growth and regeneration “An Immunohistochemical and Electron Microscopic Study”. J. Cytol. Histol. 7, 437 (2016).
Soliman, S. A., & Abdelhafeez, H. H. A new mechanism of cartilage growth in mammals “Involvement of CD117 positive undifferentiated cells in interstitial growthm. Mathews J. Cytol. Histol. 1 (2017).
Yousef, M. S., Abd-Elhafeez, H. H., Talukder, A. K., & Miyamoto, A. Ovulatory follicular fluid induces sperm phagocytosis by neutrophils, but oviductal fluid around oestrus suppresses its inflammatory effect in the buffalo oviduct in vitro. Mol. Reprod. Dev. 86(7), 835–846. https://doi.org/10.1002/mrd.23164846 (2019).
Abdel-Hakeem, S. S., Mahmoud, G. A., & Abdel-Hafeez, H. H. Evaluation and microanalysis of parasitic and bacterial agents of Egyptian fresh sushi, Salmo salar. Microsc Microanal. 3, 1–11. https://doi.org/10.1017/S143192761901506X (2019).
Soliman, S. A. & Abd Elhafeez H. H. Are C-KIT, MMP-9 and type II collagen positive undifferentiated cells involved in cartilage growth? A description of unusual interstitial type of cartilage growth. J. Cytol. Histol. 07(04) (2016).
Moustafa, F. E.-Z. A., & El-Desoky, S. M. Architecture and cellular composition of the spleen in the Japanese quail (Coturnix japonica). Microsc. Microanal 26(3), 1–10 (2020).
Mustafa, F. E.-Z. A. & Elhanbaly, R. Histological, histochemical, immunohistochemical and ultrastructural characterization of the testes of the dove. Zygote. https://doi.org/10.1017/S0967199420000477 (2020).
Matar, A. & Silverman, H. Perspectives of Egyptian research ethics committees regarding their effective functioning. J. Empir. Res. Hum. Res. Ethics 8, 32–44. https://doi.org/10.1525/jer.2013.8.1.32 (2013).
Enterline, H. & Thompson, J. The Normal Esophagus—Embryology, Structure, and Function (Springer-Verlag, 1984).
DehghaniTafti, E., Shojaei, B., Tootian, Z., Hashemnia, S. & Sheybani, M. T. Histological and histochemical study of esophagus in pheasant (Phasianus Colchicus) embryo. ASJ 14, 121–132 (2017).
Gosomji, I. J. et al. Histogenesis of the oesophagus of guinea fowl (Numida meleagris) at Prehatch and Posthatch. Scientifica 2016, 1–5 (2016).
Liebermann-Meffert, D. in Barrett’s Esophagus (eds Hugo W. Tilanus & Stephen E. A. Attwood) 17–30 (Springer, 2001).
Bevelander, G. & Hoskins, M. M. Essentials of Histology (Mosby, 1965).
Jones, H. R., De Vivo, D. C. & Darras, B. T. Neuromuscular Disorders of Infancy, Childhood, and Adolescence: A Clinician’s Approach (Butterworth-Heinemann, 2003).
Shojaei, B., Hashemnia, S. & Rad, R. Histochemical study of the esophagus in the chukar partridge (Alectoris chukar) embryo. Folia Morphol. https://doi.org/10.5603/FM.a2016.0011 (2016).
Esrefoglu, M., Taslidere, E. & Cetin, A. Development of the esophagus and stomach. Bezmialem Sci. 5, 175–182. https://doi.org/10.14235/bs.2017.811 (2018).
Ventura, A. et al. Histological description of morphogenesis of the gastroesophageal mucosa of Gallus gallus domesticus (Linnaeus, 1758). Int. J. Morphol. 31, 1331–1339. https://doi.org/10.4067/s0717-95022013000400030 (2013).
Tomar, M. Avian esophagus: a comparative microscopic study in birds with different feeding habits. (2015).
Klem, D., Brancato, C. R., Catalano, J. F. & Kuzmin, F. L. Gross morphology and general histology of the esophagus, ingluvies and proventriculus of the house sparrow (Passer domesticus). Proc. Pennsylvania Acad. Sci. 56, 141–146 (1982).
Hadi, K. & Mohamed, A. Comparative anatomical and histological study of the esophagus of local adult male and female homing pigeon (Columba livia domestica) مقارنةونسجٍةتشزٌحٍةدراسةلمزيءواألنثىالذكزًفالحماًالمحلالزاجلمالبالغ (Columba livia domestica). AL-Qadisiya J. Vet. Med. Sci. 14, 80–87 (2015).
Carlson, B. M. Human Embryology and Developmental Biology E-Book (Elsevier Health Sciences, 2008).
Slatter, D. H. Textbook of Small Animal Surgery (Saunders, 2003).
Lim, S.-S. & Low, F. N. Scanning electron microscopy of the developing alimentary canal in the chick. Am. J. Anat. 150, 149–173. https://doi.org/10.1002/aja.1001500111 (1977).
Uriona, T. J., Farmer, C. G., Dazely, J., Clayton, F. & Moore, J. Structure and function of the esophagus of the American alligator (Alligator mississippiensis). J. Exp. Biol. 208, 3047–3053. https://doi.org/10.1242/jeb.01746 (2005).
Rajabi, E. & Nabipour, A. Histological study on the oesophagus and crop in various species of wild bird. Avian Biol. Res. 2, 161–164. https://doi.org/10.3184/175815509x12474789336122 (2009).
Shiina, T. et al. A comparative histological study on the distribution of striated and smooth muscles and glands in the esophagus of wild birds and mammals. J. Vet. Med. Sci. 67, 115–117. https://doi.org/10.1292/jvms.67.115 (2005).
Abdelnaeem, A., Rady, M. & Khalil, F. Histological and histochemical studies of the esophagus and stomach in two types of birds with different feeding behaviors. Int. J. Dev. 8, 23–41. https://doi.org/10.21608/idj.2019.64030 (2019).
Akter, K., Mussa, T., Sayeed, A., Hai, M. & Uddin, M. Study on postnatal growth and development of crop and proventriculus of digestive tract of broiler. Bangladesh J. Vet. Med. 16, 7. https://doi.org/10.3329/bjvm.v16i1.37367 (2018).
Winship, D. H., Zboralske, F. F., Weber, W. N. & Soergel, K. H. Esophagus in rumination. Am. J. Physiol. Legacy Content 207, 1189–1194. https://doi.org/10.1152/ajplegacy.1964.207.6.1189 (1964).
Mahmood, H. Comparative of histological study of esophageal glands between dogs and rabbits. (2018).
Nascimento, W., Silva, G., da Silva, L., Silva, N. & Chellappa, S. Histology of the Digestive Tract of Anablepsoides urophthalmus from Brazilian Oriental Amazonia. J. Aquacult. Mar. Biol. 7, 39–42 (2018).
Awaad, A., Moawad, U. & Tawfiek, M. Comparative histomorphological and histochemical studies on the oesophagus of Nile Tilapia Oreochromis niloticus and African Catfish Clarias gariepinus. J. Histol. https://doi.org/10.1155/2014/987041 (2014).
Santos, M., Arantes, F., Pessali, T. & Dos Santos, J. Morphological, histological and histochemical analysis of the digestive tract of Trachelyopterus striatulus (Siluriformes: Auchenipteridae). Zoologia 32, 296–305. https://doi.org/10.1590/s1984-46702015000400005 (2015).
Krauss, R. S., Chihara, D. & Romer, A. I. Embracing change: Striated-for-smooth muscle replacement in esophagus development. Skelet. Muscle 6, 27–27. https://doi.org/10.1186/s13395-016-0099-1 (2016).
Al-Juboury, R. Comparative anatomical, histological and histochemical studies of the oesophagus in two different Iraqi birds (Columba palumbus and Tyto alba). (2016).
Yovchev, D. & Penchev, G. Histochemical investigation of the esophagus of the Wild bronze turkey (Meleagris gallopavo). Trakia J. Sci. 17, 308–311. https://doi.org/10.15547/tjs.2019.04.002 (2019).
Saxena, A. K. & Klimbacher, G. Comparison of esophageal submucosal glands in experimental models for esophagus tissue engineering applications. Esophagus 16, 77–84. https://doi.org/10.1007/s10388-018-0633-9 (2019).
Fagundes, K., Rotundo, M. & Mari, R. Morphological and histochemical characterization of the digestive tract of the puffer fish Sphoeroides testudineus (Linnaeus 1758) (Tetraodontiformes: Tetraodontidae). Anais Acad. Bras. Ciênc. https://doi.org/10.1590/0001-3765201620150167 (2016).
Oliveira Ribeiro, C. A. & Fanta, E. Microscopic morphology and histochemistry of the digestive system of a tropical freshwater fish Trichomycterus brasiliensis (Lütken) (Siluroidei, Trichomycteridae). Rev. Bras. Zool. 17, 953–971 (2000).
Kalhoro, H. et al. Morphological study of the gastrointestinal tract of Larimichthys crocea (Acanthopterygii: Perciformes). Zoologia 35, 1–9. https://doi.org/10.3897/zoologia.35.e25171 (2018).
Purushothaman, K. et al. Morpho-histological characterisation of the alimentary canal of an important food fish, Asian seabass (Lates calcarifer). PeerJ 4, e2377–e2377. https://doi.org/10.7717/peerj.2377 (2016).
Pedersen, A., Sørensen, C., Proctor, G. & Carpenter, G. Salivary functions in mastication, taste and textural perception, swallowing and initial digestion. Oral Dis. 24, 1399–1416. https://doi.org/10.1111/odi.12867 (2018).
Koelz, H. Gastric acid in vertebrates. Scand. J. Gastroenterol. Suppl. 193, 2–6. https://doi.org/10.3109/00365529209095998 (1992).
Beasley, D. E., Koltz, A. M., Lambert, J. E., Fierer, N. & Dunn, R. R. The Evolution of stomach acidity and its relevance to the human microbiome. PLoS ONE 10, e0134116–e0134116. https://doi.org/10.1371/journal.pone.0134116 (2015).
Reece, W. O. Dukes’ Physiology of Domestic Animals. Physiology of Smooth Muscle (Wiley, 2015).
Shehan, N. Antomical and histological study of esophagus in geese (Anser anser demesticus). Bas. J. Vet. Res 11, 13–22 (2012).
Bódi, I. In vivo identification of avian thymic dendritic cells and the compartmentalized thymic medulla PhD thesis, Semmelweis University, School of Molecular Medicine, (2016).
Summerfield, A. & McCullough, K. C. Dendritic cells in innate and adaptive immune responses against influenza virus. Viruses 1, 1022–1034. https://doi.org/10.3390/v1031022 (2009).
Patente, T. A. et al. Human dendritic cells: Their heterogeneity and clinical application potential in cancer immunotherapy. Front. Immunol. https://doi.org/10.3389/fimmu.2018.03176 (2019).
De Zeeuw, C., Hertzberg, E. & Mugnaini, E. The dendritic lamellar body: A new neuronal organelle putatively associated with dendrodendritic gap junctions. J. Neurosci. 15, 1587–1604. https://doi.org/10.1523/jneurosci.15-02-01587.1995 (1995).
Nestle, F. O., Filgueira, L., Nickoloff, B. J. & Burg, G. Human dermal dendritic cells process and present soluble protein antigens. J. Investig. Dermatol. 110, 762–766. https://doi.org/10.1046/j.1523-1747.1998.00189.x (1998).
Bani, D., Formigli, L., Gherghiceanu, M. & Faussone-Pellegrini, M. S. Telocytes as supporting cells for myocardial tissue organization in developing and adult heart. J. Cell Mol. Med. 14, 2531–2538 (2010).
Marini, M., Manetti, M., Rosa, I., Ibba-Manneschi, L. & Sgambati, E. Telocytes in human fetal skeletal muscle interstitium during early myogenesis. Acta Histochem. 120, 397–404 (2018).
Sanches, B. D. A. et al. Telocytes play a key role in prostate tissue organisation during the gland morphogenesis. J. Cell Mol. Med. 21, 3309–3321. https://doi.org/10.1111/jcmm.13234 (2017).
Zheng, Y. et al. Genetic comparison of mouse lung telocytes with mesenchymal stem cells and fibroblasts. J. Cell Mol. Med. 17, 567–577. https://doi.org/10.1111/jcmm.12052 (2013).
Bei, Y., Wang, F., Yang, C. & Xiao, J. Telocytes in regenerative medicine. J. Cell Mol. Med. 19, 1441–1454. https://doi.org/10.1111/jcmm.12594 (2015).
Sidney, L. E., Branch, M. J., Dunphy, S. E., Dua, H. S. & Hopkinson, A. Concise review: Evidence for CD34 as a common marker for diverse progenitors. Stem Cells 32, 1380–1389. https://doi.org/10.1002/stem.1661 (2014).
Nielsen, J. S. & McNagny, K. M. Novel functions of the CD34 family. J. Cell Sci. 121, 3683–3692. https://doi.org/10.1242/jcs.037507 (2008).
Wang, L. et al. A small-molecule/cytokine combination enhances hematopoietic stem cell proliferation via inhibition of cell differentiation. Stem Cell Res. Ther. 8, 169. https://doi.org/10.1186/s13287-017-0625-z (2017).
Madihally, S. V., Flake, A. W. & Matthew, H. W. T. Maintenance of CD34 expression during proliferation of CD34+ cord blood cells on glycosaminoglycan surfaces. Stem Cells 17, 295–305. https://doi.org/10.1002/stem.170295 (1999).
Adair, T. H. & Montani, J. P. in Chapter 1, Overview of Angiogenesis (2010).
Iruela-Arispe, M. L. & Zovein, A. in Fetal and Neonatal Physiology (Fifth Edition) (eds Richard A. Polin et al.) 85–89 (Elsevier, 2017).
Bojin, F. M. et al. Telocytes within human skeletal muscle stem cell niche. J. Cell Mol. Med. 15, 2269–2272. https://doi.org/10.1111/j.1582-4934.2011.01386.x (2011).
Ito, T. K., Ishii, G., Chiba, H. & Ochiai, A. The VEGF angiogenic switch of broblasts is regulated by MMP-7 from cancer cells. Oncogene 26, 7194–7203. https://doi.org/10.1038/sj.onc.1210535 (2007).
Pammer, J. et al. Vascular endothelial growth factor is constitutively expressed in normal human salivary glands and is secreted in the saliva of healthy individuals. J. Pathol. 186, 186–191 (1998).
Keswani, S. G. et al. Role of salivary vascular endothelial growth factor (VEGF) in palatal mucosal wound healing. Wound Repair. Regen. 21, 554–562. https://doi.org/10.1111/wrr.12065 (2013).
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The work was equally distributed between authors; S.A.S., and F.M., including designed the research study, the analysis. S.A.S. interpreted the data, arranged the images and wrote the paper. Dr S.A.S. drew Fig. 15. All authors have read and approved the final version of the manuscript.
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Soliman, S.A., Madkour, F.A. Developmental events and cellular changes occurred during esophageal development of quail embryos. Sci Rep 11, 7257 (2021). https://doi.org/10.1038/s41598-021-86503-9
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DOI: https://doi.org/10.1038/s41598-021-86503-9
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