Open Veterinary Journal, (2026), Vol. 16(4): 2488-2504

Research Article

10.5455/OVJ.2026.v16.i4.49

Variance expression of MUC17 and mucin histochemistry in the gastrointestinal tract of the desert-adapted Euphrates jerboa (Jaculus euphraticus)

Raed Gahat Mehjal¹, Manar Mousa Alhussein², Abdulrazzaq B. Kadhim^1*, Hazem Almhanna¹, Nabeel Abd Murad Al-Mamoori¹ and Haneen Abdul Ameer Abbas¹

¹Department of Anatomy and Histology, College of Veterinary Medicine, University of Al-Qadisiyah, Al-Diwaniyah City, Iraq

²Department of Pathological Analysis, College of Science, University of Al-Qadisiyah, Al-Diwaniyah City, Iraq

*Corresponding Author: Abdulrazzaq B. Kadhim. Department of Anatomy and Histology, College of Veterinary Medicine, University of Al-Qadisiyah, Al-Diwaniyah City, Iraq. Email: Abdulrazzaq.alrabei [at] qu.edu.iq

Submitted: 29/11/2025 Revised: 17/03/2026 Accepted: 29/03/2026 Published: 30/04/2026

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ABSTRACT

Background: Euphrates jerboa is a desert-adapted animal that survives in extreme arid conditions in Iraq and other countries. This might be due to specialized gastrointestinal (GI) adaptations that allow this animal to survive hard conditions.

Aim: This study aimed to investigate the morphological, histochemical, and molecular features of the esophagus, stomach, and duodenum.

Methods: Traditional histomorphology, histochemistry, and real-time polymerase chain reaction targeting the membrane-bound mucin (MUC17) gene were conducted.

Results: Morphometry revealed a short esophagus, a J-shaped stomach, and an elongated duodenum, indicating adaptations for processing a dry, fibrous diet. Histochemistry results exhibited region-specific mucin scattering. In particular, neutral mucins dominated gastric surface cells of the stomach and duodenal Brunner’s glands of the duodenum, whereas acidic mucins were abundant in the esophageal submucosal glands of the esophagus and gastric neck cells of the stomach. Real-time quantitative polymerase chain reaction revealed higher MUC17 gene expression in the stomach than in the esophagus and duodenum (p=0.009), representing a stomach-specific role for this membrane-bound mucin.

Conclusion: The results indicated that in the GI of jerboa, more mucin secretion and different expression status of MUC17 were recruited to reinforce its mucosal barrier combined with lubrication and water conservation for jerboa against extreme aridity conditions in deserts. This integrated strategy uncovers major evolutionary adaptations involved in aridity survival and establishes the jerboa as a new model to investigate mucosal resilience and mucin-related diseases.

Keywords: Desert adaptation, Gastrointestinal tract, MUC17, Mucosal defense, RT-Qpcr.

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Introduction

The gastrointestinal (GI) tract is an important system for animals in which nutrients are absorbed, mucosa is protected, and the immune defense mechanism against different organisms such as parasites, bacteria, and viruses. In brief, jerboas are small, bipedal, rodent-like animals that live in arid and semiarid areas where life is a challenge: water is limited, temperatures are very high, and the diet consists of fibrous vegetal matter with very little moisture (Withers et al., 2016). Such a tough life demands special physiological and molecular adaptations so that the animal can maintain proper digestion and conserve water (Schmidt-Nielsen, 1997).

Many studies have focused on animal models and have asserted that the esophagus of rodents has four layers: the mucosa (stratified squamous epithelium, lamina propria, and muscularis), submucosa, muscularis, and adventitia (Uehara et al., 2018; Al-Taai, 2022). Consequently, the epithelium consisted of non-keratinized African Giant Rats, and the lamina propria was mostly rich with collagen/elastic fibers, but the muscularis longitudinal smooth muscle (absent in African Giant Rats. In addition, the submucosa displays a weak periodic acid-Schiff’s (PAS) reaction, and the muscularis reacts with less intensity to PAS (Nzalak et al., 2010; Hassani, 2021).

In addition, rodent stomachs are similar to those of other mammals, comprising the mucosa, submucosa, muscularis externa, and serosa. The stomach has a unicular, C-shaped shape and lacks external demarcation between regions. However, the cardiac region lines were stratified by squamous epithelium with mucous glands. The fundic region illustrates thick mucosa compared with other regions with parietal/chief cells. Similarly, the pyloric region shows branched glands. Muscularis extends to the esophagus and contains both skeletal/smooth muscle in the cardiac region and smooth muscle elsewhere (Wattel, 1978; Krauss et al., 2016; Elsayed and El-Gammal, 2024).

The rodent duodenum comprises four layers: the mucosa, submucosa, muscularis externa, and serosa. The mucosa comprises absorptive enterocytes with microvilli, goblet cells, and Paneth cells. The submucosa has well-developed Brunner’s glands that secrete alkaline mucus, which is more pronounced than that in humans. The muscularis externa has smooth (circular and longitudinal) muscles with the myenteric plexus sandwiched between them. Rodents show leaf-shaped villi, fast turnover of epithelium (3–5 days), and apparently stronger mucosal protection, all of which are adaptations to their abrasive diet. These histological features indicate the specialized digestive and protective functions of rodents (Treuting et al., 2017; Nakatsuji et al., 2018; Imeer et al., 2023).

A mucin is a massive glycoprotein secreted from epithelial cells and, hence, constitutes furiously essential components of mucous hanging off on the GI tract lining as gel or epithelial mucins for epithelium or lumen of organs. Membrane-bound mucin (MUC17) is a membrane-associated mucin involved in sustaining epithelial integrity and modulating cellular signaling in the gut. The expression and roles of MUC17 have been studied in conventional laboratory rodents and humans but not in desert-adapted species such as the Euphoretic Jerboa. Molecular processes coordinating these adaptations remain obscure, especially the coordinated action of mucins, such as MUC17, in mucosal defense and nutrient absorption (Gum Jr et al., 2002; Luu et al., 2010; Jäverfelt, 2023; Fan et al., 2025).

Therefore, this study aimed to baseline explore tissue-level adaptations in the GI tract and gene expression patterns of jerboa using histomorphological study and quantitative PCR (qPCR) techniques. Finally, this study aims to investigate the functions of species-specific and related recognition type mucins to reveal novel evolutionary adaptations that may feed into biomedicine and ecology.

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Materials and Methods

Animals

Healthy adult jerboas (euphoretic jerboa) weighing between 30 and 50 g were collected from the northeastern region of Al-Qadisiyah Province, weighting between 30 and 50 g. The animals were inspected in the laboratory, and healthy animals were selected for this study. All experimental procedures followed strict ethical guidelines for animal research and were approved by the relevant authorities of the College of Veterinary Medicine/University of Al Qadisiyah. Before the experiments, the Jerboas were acclimated for at least 2 weeks in a controlled environment with a 12-hour light/dark cycle (8 am–8 pm) and a constant temperature of 23°C ± 0.6°C. They had unlimited suitable food and water before killing the animals.

Collection of the samples

Specimens were collected from the alimentary canal, including the esophagus, stomach, and duodenum. Organs were dissected for morphological study, and length, width, thickness, and weight were measured. Specimens for histology were dissected and immediately washed and fixed using 10% formalin solution for 24–48 hour. Moreover, specimens from the same regions were collected for gene expression study and saved directly in Trizole (Scientific researcher co. Ltd, Iraq).

Histomorphological techniques

For histological examinations, the esophagus, stomach, and duodenum specimens were collected immediately after dissection, washed using distilled water, and fixed for 24–48 hour in 10% formalin solution. The samples were then dehydrated in ascending grades of ethyl alcohol, cleared in xylene, and embedded in paraffin wax. The sections cut at 5 µm level. All sections were routinely stained with hematoxylin and eosin to detect the general structure of the tissue, and Masson trichrome, Vierhoff’s PAS, and Alcian blue (PH.2.5) stains were used to determine the special contents of the tissue, as described previously (18). For quantified analysis, epithelial cell populations were quantified and characterized for three histological regions, including the cardiac, fundic, and pyloric regions of the stomach. All different sections were examined and images were captured using a light microscope (Olympic^mt, Japan) at a magnification of 4×,10×,20× and 40×.

mRNA expression

Total Ribonucleic acid (RNA) was isolated from the esophagus, stomach, and duodenum tissue of jerboa using the Accuzol® reagent system (Bioneer, Korea). The tissue samples (150 mg) were homogenized in 1.5 ml microcentrifuge tubes, had 200 μl chloroform (Sigma-Aldrich, USA) was added, and then vigorously vortexed. After incubation on ice for 5 minutes, the samples were centrifuged at 10,000 × g for 15 minutes at 4°C. The aqueous phase was mixed with an equal volume of isopropanol (Sigma-Aldrich, USA) and held at 4°C for 10 minutes. After another 10 minutes of centrifugation (10,000 × g, 4°C), the RNA pellets were washed with 1 ml of 80% ethanol (Sigma-Aldrich, USA). After the final centrifugation, the pellets were left to air-dry and then suspended in 50 μl of DEPC-treated water (Sigma-Aldrich, USA) and stored at −20°C. The RNA concentration and purity were determined using Nanodrops (Thermo Fisher Scientific, USA).

cDNA synthesis

DNase I treatment was performed on the extracted RNA to eliminate possible contamination of genomic DNA (Promega, USA). First, cDNA synthesis was then performed, employing DiaStar™ OneStep RT-PCR Kit (China) following the manufacturer’s instructions. The synthesized cDNA was equally concentrated and stored at −20°C until further analysis.

Quantitative-PCR analysis

MUC17 expression levels were detected through qPCR using SYBR green on a BioRad CFX96 machine (USA). Gene-specific primers (designed by Scientific Researcher Co. Ltd, Iraq) of MUC17 (XM_045142857.1) and Euphoretic Jerboa GAPDH (reference gene) (XM_015873412.2) are listed in Table 1. The qPCR protocol consisted of: Reverse transcription: 50°C for 60 minutes (one cycle), polymerase chain reaction (PCR) amplification: 95°C for 20 seconds, followed by 40 cycles of 95°C for 20 seconds and 60°C for 30 seconds. Melt Curve analysis arranged 60°C–95°C with increments of 0.5°C. The reactions used AccuPowerTM 2X Green Star qPCR master mix (Bioneer, Korea), with fluorescence detection at each extension step. Data were normalized against GAPDH expression via the 2-ΔΔCt method (Livak and Schmittgen, (2001).

Table 1. Primer sequence used for MUC17 and GAPDH detection.

Statistical analysis

Morphological measurements were collected, and the mean + standard error of the esophagus, stomach, and duodenum, including weight, relative weight, length, width, and thickness, were measured and analyzed using the t-test. The average mucin quantities were then investigated. Raw data on MUC17 gene expression in identical tissues were collected using Real-time quantitative polymerase chain reaction (RT-qPCR) and evaluated using 2^ΔCT (19). Statistical analysis was conducted using the Statistical Package for the Social Sciences. A two-way analysis of variance (ANOVA) was used to examine mucin quantities, whereas a one-way ANOVA was used for RT-qPCR. P-values 0.05 indicate significance.

Ethical approval

The project was approved by the College of Veterinary Medicine, University of Al-Qadisiyah (1890-22-11-2022).

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Results

Morphometric results

This study characterized and described the morphological and gross structures of the esophagus, stomach, and duodenum. Therefore, the esophagus was a tube-shaped organ with a multi-layered muscular membrane that appeared glossy and fluctuated in color from white to pink. It was extended from the pharynx to the cardia of the stomach. This organ showed three portions: the cervical, thoracic, and abdominal portions (Fig. 1). The cervical part runs the full length of the neck, starting at the lower end of the pharynx esophagus and ending at the point where the first ribs meet the right thoracic inlet parallel to the borders of the left and right common carotid arteries on both sides. The thoracic portion started at the thoracic inlet, parallel to the caudal vena cava, and acted as a route through the diaphragm before entering the esophageal hiatus. The short part of the esophagus was the abdominal portion and extended as an esophageal hiatus to consist of the of the gastroesophageal junction to the stomach, and it formed an impression groove on the parietal surface of the left lobe of the liver (Fig. 1). A thicker area of muscle fibers encircled the esophagus base, forming a circular muscle as the esophageal sphincter. The mean (weight, relative weight, length, width, and thickness) values of the esophagus were accounted for 0.02 + 0.01 am., 0.014 + 0.00 am., and 3 + 0.07 cm. 0.1 + 0.0 mm and 0.1 + 0.0 mm (Table 1).

Fig. 1. A. Morphological section of the elementary canal of the jerboa shows the course of the esophagus (green arrow) from the pharynx opening (blue arrow) extending between the tracheal bifurcation (white arrow) and the thoracic part (yellow arrow) penetrating the diaphragm (brown arrow) to the stomach, and the liver (red arrow) covering the stomach. The duodenum (black arrow). B. Cervical (blue line) and thoracic (yellow line) parts.

The stomach was also directed vertically within the abdominal cavity, connected between the esophagus and the small intestine. The retro-diaphragm and retro-hepatic areas were positioned closer to the left side of the midline than to the right side. the appeared J-shaped with uniform gray color and has two openings (the cardiac and pyloric), two surfaces (the parietal and visceral), two curves (the lesser and greater curvatures), and two ends (Fig. 2A). Internally, the stomach was unilocular, exhibiting a main fold between the cardiac and fundic regions as well as small folds extending inside the stomach regions. The stomach displayed three main parts: the cardiac area, which was thicker and rougher and surrounded the esophagus at the top; the fundic area, which included the fundus and body and made up most of the stomach; and the pyloric area, which had a funnel-shaped connection to the duodenum. The lesser curvature was quite short, linking to the small intestine via the omentum, whereas the greater curvature was broad and associated with the spleen, extending slightly toward the back (Fig. 3A). In the pyloric region, the pyloric antrum connects to the main part of the stomach and leads into the pyloric canal, which goes toward the duodenum at the pyloric sphincter (Fig. 3B). The mean (weight, relative weight, length, width, and thickness) of the stomach parameters were accounted for 0.66 + 0.04 gm and 0.36 + 0.007 gm, 1.62 + 0.08 cm. 0.64 + 0.05 mm and 0.46 + 0.02 mm (Table 2).

Fig. 2. A. Morphological section of the esophagus, stomach, and duodenum of jerboa shows: the esophagus parts are cervical (blue arrow), thoracic (yellow arrow), abdominal (green arrow) (blue line), and thoracic (yellow line). Stomach (black arrow) and duodenum (black arrow). B. Duodenal length (blue line) and spleen (yellow arrow).

Fig. 3. A. Macroscopic section of the internal surface of stomach of jerboa shows: the esophagus opening (blue arrow), duodenal opening (red arrow) and main fold (yellow arrow), no glandular part (blue star), and glandular part (black arrow). B. External appearance of the stomach: esophagus (white arrow), duodenum stomach junction (orange arrow), and stomach corpus (green arrow).

Table 2. Morphometric parameter analysis of the esophagus, stomach, and duodenum of jerboa.

In addition, the duodenum was tube-like and connected to the pyloric opening of the stomach, forming three main parts: superior, descending, and ascending, extending to the sigmoid flexure that disconnected between the duodenum and jejunum (Fig. 2B). The mean (weight, relative weight, length, width, and thickness) of the duodenum measurements were accounted for 0.38 + 0.03 gm, 0.22 + 0.00 gm, 7.6 + 0.37 cm, 0.4 + 0.0, and 0.16 + 0.024 mm (Table 3).

Table 3. This table shows the quantitative comparison of relative MUC17 gene expression patterns across the esophagus, stomach, and duodenum.

Histological results

Histological description of the esophagus, stomach, and duodenum of the Euphrates jerboa using H- and E-stained sections, and the special histological content of each organ. Therefore, the esophageal wall was structured in four distinct layers, including the mucosa, submucosa, muscularis externa, and adventitia/serosa (Fig. 4). The mucosa was lined with non-keratinized stratified squamous epithelium, which contains a clear basal cell layer and a thin lamina propria with dispersed lymphocytes based on longitudinal muscularis mucosae (Fig. 5). The submucosa displayed dense irregular connective tissue and prominent esophageal glands (mucous type). The submucosa was based on the muscularis externa and consisted of two pattern muscles arranged in the inner circular and outer longitudinal layers (Figs. 6 and 7). In addition, the proximal part comprised a mix of skeletal and smooth muscle fibers, whereas the distal part transitioned to purely smooth muscle (Fig. 8). Finally, the adventitia/serosa was the outermost layer consisting of fibroblastic connective tissue rich in lymphocytes, blood vessels, and adipose tissue (Fig. 8).

Fig. 4. Histological section of the cervical part of the esophagus showing the mucosa (yellow arrow), ...(upper blue arrow), lamina propria (green star), submucosa (red arrow), striated muscles (red star), and adventitia (lower blue arrow). H&E x100.

Fig. 5. Histological section of the esophagus shows the following mucosa cell layers: keratinized (black arrow), granulosa cells (white arrow), spinous cells (yellow arrow), and basal cells (red arrow) mucosa muscularis (blue arrow). H&E x400.

Fig. 6. Histological section of the esophagus shows: mucosa cells layers (red arrow) keratinized (black arrow), mucosa muscularis (blue arrow), submucosa (yellow arrow), muscular layers (yellow line) and serosa (black star). H&E ×100.

Fig.7. Histological section of the esophagus shows: three layers of muscularis: Muscularis externa (yellow line), Muscularis intermediate (black line), and Muscularis interna (red line) .H&E x400.

Fig. 8. Histological section of the esophagus shows: proximal part (A) and distal (B) parts. H&E x400.

Comparison of the histological staining of the esophagus of Euphrates jerboa with PAS and Alcian blue reveals key differences in the target substances and their localization. The Alcian blue stain appears as a teal or greenish-blue color and specifically targets acidic mucosubstances, with strong staining localized to the Submucosal glands. In contrast, the PAS stain appears as a magenta or pink-purple color and stains mucopolysaccharides. PAS staining is localized to mucus-producing cells within both the submucosa and epithelial layer. Both stains show a strong presence of mucin, but they highlight different chemical components and have slightly different distribution patterns within the esophageal tissue (Fig. 9).

Fig. 9. Histochemical section of the esophagus shows: reaction the esophagus layers with H&E (A, D), PAS (B), Alcian blue (C), Massontrichom (E), and Verhoffer stain (F). x100.

Similarly, the stomach wall was composed of the innermost histological layers: mucosa, submucosa, muscularis externa, and serosa, using H and E staining. The mucosa contained gastric pits and glands and was characterized by three specialized regions, including the cardiac, fundic, and pyloric regions (Fig. 10). Consequently, the cardiac region is lined by simple columnar epithelium with shallow gastric pits and branched tubular cardiac glands that primarily secrete mucus. The fundic region was shoed deep into the gastric pits and was packed with dense parietal cells (acidophilic cytoplasm) and chief cells (rich with basophilic zymogen granules) (Fig. 11). In addition, the pyloric region was distinguished by very deep gastric pits and coiled, branched pyloric glands rich in mucus-secreting cells (Fig. 12).

Fig. 10. Histological section of the cardiac region of stomach shows: epithelia no keratinized stratified squamous (blue arrow), lamina propria (black star), submucosal tumor (yellow arrow), muscularis externa (white line), and serosa (black arrow). H&E x100.

Fig. 11. Histological section of the fundic region of the stomach shows the following: simple columnar epithelia (blue arrow), mucosa muscularis (yellow arrow), submucosal tumor (black arrow), externa muscularis (blue line), and serosa (green arrow). H&E x100.

Fig. 12. Histological section of the fundic region of the stomach shows the following: surface mucosa cells (blue arrow), mucus neck cells (yellow arrow), parietal cells (black arrow), chief cells (red arrow), and endocrine cells (white arrow). H&E x100.

The submucosa supported the mucosa and contained blood vessels, lymphatics, and nerves. The muscularis externa was a compact and thick layer with different directions of smooth muscle fibers. The pattern of the muscularis externa displayed three layers of muscle fiber directions: inner oblique, middle circular, and outer longitudinal, with the myenteric (Auerbach’s) plexus observable between them.

Finally, the outermost layer of the stomach was the serosa, which is a protective covering comprising fibroblastic connective tissue rich with lymphocytes, blood vessels, and adipose tissue (Fig. 13).

Fig. 13. Histological section of the pyloric region of the stomach shows the following: mucosa (yellow star), submucosa (blue star), muscularis externa (black star), and pyloric gland (blue arrow). H&E x100, 400.

Using the special stains, histological sections of the stomach stained with combined PAS and Alcian blue revealed distinct mucosubstance distribution. The gastric mucosa exhibited characteristic glandular structures. Alcian blue (bluish green), indicating acidic mucosubstances, was strongly localized to the mucus neck cells of the gastric and pyloric glands. The pinkish-purple pigment identifying neutral mucosubstances was predominantly observed in the surface mucous cells. Both stains exhibited robust intensity in their respective areas. Areas with purplish coloration indicated the presence of both acidic and neutral mucosubstances (Fig. 14).

Fig. 14. Histochemical section of the stomach shows: reaction the esophagus layers with PAS (B), Alcian blue (C), Massontrichom (E), and Vierhoff's stain (F) x100. H&E staining A and D.

In contrast, the duodenum demonstrated the typical intestinal wall structure of mammals: mucosa, submucosa, muscularis externa, and serosa. Therefore, the mucosa was lined with a tall simple columnar epithelium with a brush border and abundant scattered goblet cells. Moreover, the Lieberkühn crypts extended toward the muscularis mucosae, and the villi were elongated and projected from epithelial cells and directed into the lumen. This layer is based on the submucosa, which consists of dense connective tissue with a rich vascular network and is notable for its characteristic Brunner’s glands, which are branched tubular mucous glands. The muscularis externa consisted of a thicker inner circular layer and an outer longitudinal layer, with a regular myenteric plexus. Finally, the serosa consisted of a thin mesothelial covering with underlying connective tissue.

Our study found that histochemical staining of the duodenum with PAS and Alcian blue (AB) presented distinct mucin distribution patterns in the duodenal mucosa of Euphrates jerboa. Therefore, the Brunner’s glands displayed strong PAS reactivity, signifying abundant neutral mucins, with moderate AB staining intensity demonstrating concurrent acidic mucin production. In addition, goblet cells revealed variable staining intensities, with predominant PAS positivity compared with weaker AB reactivity. In addition, the muscularis mucosae, enterocyte glycocalyx, and lamina propria exhibited persistent PAS positivity (+), whereas AB staining was insignificant in these structures.

Moreover, we detected distributed collagen and elastic fibers in different locations of the esophagus, stomach, and duodenum using Masson trichrome and Verhoeff staining. In the histological parts of the esophagus, collagen fibers were excessively detected in the epithelium and submucosa and less in the muscularis tunica. Elastic fibers were scattered and consisted of the epithelium and tunica muscularis, with less in the submucosa and absent in the lamina propria (Fig. 15). In the stomach, a high distribution of collagen fibers is observed in the lamina propria and tunica muscularis and less in the submucosa and tunica adventitia. Elastic fibers were clearly interspersed in most stomach layers, including the tunica mucosa and tunica muscularis, and in the submucosa and tunica adventitia (Fig. 16). Similarly, duodenal tissue showed high collagen fibers in the lamina propria, whereas more elastic fibers were found in the epithelium, submucosa, and tunica muscularis, and less in the lamina propria and tunica adventitia (Fig. 17).

Fig. 15. The histological section of the duodenum shows: mucosa (blue arrow), crypt of the librkahan (yellow arrow), muscularis (red arrow), submucosa (black arrow), externa (white arrow), and serosa (green arrow). H&E x100, 400.

Fig. 16. Histological section of the duodenum shows: duodenal gland (yellow arrow), mucosa muscularis (red arrow), and Brunner’s glands (green arrow). H&E x400, 400.

Fig. 17. Histochemical section of the duodenum shows: reaction the esophagus layers with PAS (B), Alcian blue (C), Massontrichom (E), and Vierhoff's stain (F) x100. H&E staining A and D.

RT-qPCR results

Our results showed that MUC17 gene expression (relative to GAPDH) differed significantly among the stomach, duodenum, and esophagus of Euphrates jerboa.

The RT-qPCR assay showed high specificity and reliability of MUC17 expression measurements for amplification plots and melt curves. Therefore, the amplification curves (RFU vs. cycles) for MUC17 and the reference gene GAPDH demonstrated clear exponential amplification for both genes, and RFU values stabilize after ~30 cycles, indicating efficient PCR and sufficient product accumulation. In addition, no primer-dimer reaction suggested minimal nonspecific amplification (Fig. 18).

Fig. 18. The raw RT-qPCR amplification plot corresponding to the GAPDH gene, the PCR-qRT reaction confirms the quality and consistency of RT-qPCR data for the reference gene (GAPDH) across all samples. High specificity and efficient amplification of the reference gene are crucial for accurate normalization and reliable quantification of the target gene (MUC17) during subsequent analysis.

Interestingly, the stomach samples exhibited earlier Ct values (~25 cycles for MUC17 vs. ~29 for GAPDH), aligning with higher MUC17 expression in the esophagus and duodenum, which supported the reproducibility and consistency of the results. In addition, the melt peaks (d(RFU)/dT vs. temperature) confirm amplicon specificity via a single sharp peak, and Tm consistency across replicates validates target-specific amplification. (Fig. 18). Our data presented the region-specific expression patterns of MUC17 across the stomach, duodenum, and esophagus of Euphrates jerboa.

Therefore, the stomach demonstrated significantly higher MUC17 expression than the duodenum (p=0.009) and esophagus (p=0.009). The duodenum showed no significant difference (p=0.998) with the esophagus, as shown in Fig. 18 and Table 4.

Table 4. This table shows the statistical analysis of the MUC17 gene expression pattern across the esophagus, stomach, and duodenum.

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Discussion

In the current study, a comprehensive analysis of the morphological, histochemical, and molecular adaptations of the gastrointestinal tract in the desert-adapted Euphrates jerboa (Euphoretic Jerboa) was performed, with particular focus on the gene expression and distribution of mucins, particularly MUC17. Our investigation revealed important region-specific variations in the histological structures, mucins, and MUC17 expression that may support this species’ significant tolerance and living in an arid, high-fiber environment.

Accordingly, the morphometric results showed that the GI tract structure in jerboas, including the esophagus, stomach, and duodenum, was relatively short, with a very distinct, and had a single Type J bend of the stomach, which is generally consistent with rodent anatomy (Treuting et al., 2017; Hryn et al., 2018). Similarly, the esophageal portions (cervical, thoracic, and abdominal) and the inner folding of the stomach (Uehara et al., 2018; Alshreefy, 2024) were useful in the structural contextualization of the functional histology and physiology of jerboas. Importantly, the larger length of the duodenum compared with that of the stomach might be an evolutionary adaptation for maximizing nutrient and water absorption from a dry, fibrous diet, and this may be permitting a longer retention time of chyme for efficient digestion in an air-conditioned environment (Mackie, 2002; Alshreefy, 2024).

Furthermore, histochemical staining with PAS and Alcian Blue stains revealed region-specific mucin distribution patterns for each organ. Therefore, the strong presence of both neutral and acidic mucosubstances inside the submucosal glands and epithelial layer in the esophagus suggests a dual protective mechanism and physical barrier lined epithelium from the external environment inside the esophagus. In addition, the presence of a mucous coating is crucial for lubricating the esophagus of dry, abrasive food in the desert, thereby reducing mechanical damage and decreasing water loss during swallowing. This finding agrees with studies in other rodents that have similar dietary challenges; however, the intensity and configuration of the dietary challenge were finely recorded in the same desert species (Yandrapu and Sarosiek, 2015; Lin et al., 2020; Imeer et al., 2023).

Furthermore, the stomach demonstrated a clear functional zonation and co-localization of neutral mucins in surface mucous cells, whereas acidic mucins were found in the mucus neck and pyloric glands, confirming that they play a major role in the multi-layered defensive barrier of the stomach. The epithelial mucus, which is rich in neutral mucins, likely serves as a principal physical barrier preventing autodigestion and mechanical damage to the epithelium of the stomach (Slomiany and Slomiany, 1991; Marriott and Gregory, 2024). In contrast, the acidic mucins in the glandular regions could contribute to maintaining the optimal pH microenvironment for enzymatic activity and additional protection of the gastric epithelium of the stomach. The histology of the fundic region was rich in parietal producing Hydrochloric acid and chief cells secreting enzymes, indicating its main role in chemical digestion, which is essential to break down tough plant matter in deserts (Gomez-Santos et al., 2021; Sary et al., 2025).

The duodenum displayed a typical adaptation for nutrient absorption with elongated villi and a clear brush border. In particular, Brunner’s glands were highly stained with PAS, indicating a secretion rich in neutral mucins. This alkaline mucus is central to neutralizing the acidic chyme coming from the stomach and protecting the delicate duodenal mucosa while setting the proper pH for pancreatic enzymes. The patterned staining of the goblet cells provides further evidence of a variable mucus layer involved in lubrication and protection of the intestinal tract (Konturek et al., 2004; Desseyn et al., 2008).

The greater finding in this study must have been that membrane-bound mucin MUC17 was differentially expressed. Therefore, quantitative PCR detected a significantly higher expression of MUC17 mRNA in the stomach than in the esophagus or duodenum. Such differential expression points toward a critical stomach-specific role of MUC17 in the jerboa. As a membrane-bound mucin, MUC17 provides more lubrication and maintains epithelial integrity, forming the glycocalyx, and regulating cell signaling and proliferation at the mucosal surface (Gum Jr et al., 1994; Desseyn et al., 2008).

The highest expression of MUC17 in the stomach suggests a strengthened defense mechanism at the cellular level. Therefore, this is an important adaptation for an animal eating a dry, roughage-type diet because its gastric epithelium is constantly mechanically and chemically stressed, and this could exposure to physical damage and ulcers during the quick digestion process with water limitations.

According to our observations, the reported non-significance of MUC17 expression between the duodenum and esophagus, despite their differing histological mucin patterns, explains the complex and multifaceted nature of mucosal protection. While the duodenum depends largely on mucins secreted from Brunner’s glands for majority protection, the esophagus and much of the duodenum use this MUC17 almost at equal levels; therefore, there would have to be much more stomach investment into this particular mechanism.

The observed complementary distribution of neutral and acidic mucins may involve a dual protective measure that increases jerboa mucosal defense against varying stimuli in their high-stress GI tract. Neutral mucins, which are particularly prominent in the brunner’s glands of the stomach and duodenum, are designed to form a viscoelastic layer of mucus that forms a protective barrier on the epithelium against the damaging effects of gastric acids and digestive enzymes. Under highly luminal conditions, the barrier provides a slower rate of diffusion for hydrogen ions, traps bicarbonate ions to neutralize the acidity of the mucus barrier and reduces substrate autodigestion in the epithelium (Desseyn et al., 2008). Acidic mucins from the esophageal glands are negatively charged due to sialylation and sulfonation, which stimulates increased hydration and more vigorous interactions with irritants and microbes present in the lumen, thereby enhancing the chemically resilient barrier and protecting the pre-epithelial barrier from enzymatic damage and microbial adhesion. Overall, this specialized selection of mucin provides an integrated system of defense on the various segments of the gut, providing protection from the permeable and abrasive elements of the diet that are characteristic of xeric-adapted species (Liu et al., 2025).

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Conclusion

The interdisciplinary approach in pursuing evidence revealed that Euphrates jerboa has integrated and concerted adaptations manifesting at morphological, histochemical, and molecular levels to survive in an extreme desert environment. Differentiated mucin secretion and MUC17 gene expression upregulation in the stomach suggest a highly evolved system for mucosal protection and water conservation. Thus, these observations enrich the evolutionary biology of desert mammals and propose the jerboa as a new model for mucosal resilience research. Additionally, it pushes researchers to get more understanding behind its resilient mucosal defense system, especially the role of MUC17, it may open new biomedical pathways for the study of human diseases with mucosal breakdown, such as gastric ulcers, inflammatory bowel disease, and xerostomia.

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Acknowledgment

The authors thank the College of Veterinary Medicine, University of Al-Qadisiyah, for their support in this study.

Conflict of interest

The authors declare no conflicts of interest.

Funding

The authors have self-funded the study. No external funding source is available.

Authors’ contributions

All authors have participated in the study.

Data availability

Data are available when requested by the corresponding author.

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References

Alshreefy, M.N.A. 2024. Histomorphological and histochemical study of esophagus and stomach in adult guinea pigs (Cavia porcellus). Wasit. J. Pure. Sci. 3(2), 306–314; doi:10.31185/wjps.412

Al-Taai, S.A. 2022. Esophagus. In Pharynx—The incredible rendezvous sites of gas, liquid and solid. London, United Kingdom: IntechOpen.

Desseyn, J.L., Tetaert, D. and Gouyer, V. 2008. Architecture of the large membrane-bound mucins. Gene 410(2), 215–222; doi:10.1016/j.gene.2007.12.014

Elsayed, A.H. and El-Gammal, S.M. 2024. Comparative study on the gross anatomy of some abdominal organs in albino rat and albino mouse. Alex. J. Vet. Sci. 80(1), 1–17; doi: 10.5455/ajvs.183386

Fan, F., Guo, R., Pan, K., Xu, H. and Chu, X. 2025. Mucus and mucin: changes in the mucus barrier in disease states. Tissue Barriers 13, 2499752; doi:10.1080/21688370.2025.2499752

Gómez-Santos, L., Alonso, E., Crende, O., Ibarretxe, G., Madrid, J.F. and Sáez, F.J. 2021. Identification of sugar moieties in chief cells of the rat fundic gastric glands. Anat. Sci. Int. 96(2), 221–230; doi:10.1007/s12565-020-00578-4

Gum, J.R., Crawley, S.C., Hicks, J.W., Szymkowski, D.E. and Kim, Y.S. 2002. MUC17, a novel membrane-tethered mucin. Biochem. Biophys. Res. Commun. 291(3), 466–475; doi:10.1016/bbrc.2002.6475

Gum, Jr, J.R., Hicks, J.W., Toribara, N.W., Siddiki, B., Kim, Y.S. 1994. Molecular cloning of human intestinal mucin (MUC2) cDNA. J. Biol. Chem. 269(4), 2440–2446.

Hassani, M. 2021. Histological and histochemical examination of mucous cells in esophagus and stomach of Rattus norvegicus. Iran. J. War. Public. Health. 13(4), 289–304.

Hryn, V.H., Kostylenko, Y.P., Yushchenko, Y.P., Lavrenko, A.V. and Ryabushko, O.B. 2018. General comparative anatomy of human and white rat digestive systems: a bibliographic analysis. J. Title Not Provided 71(8), 1599–1602.

Imeer, A.T.A., Alamir, H.T.A., Albassam, T.A. and Kadhum, A.A.H. 2023. Histological and histochemical developmental study of the duodenal glands in rabbit and mice during different ages. J. Nat. Sci. Biol. Med. 14(2), 246–252.

Jäverfelt, S. 2023. The functional role of the MUC17-based glycocalyx. Gothenburg, Sweden: Univ Gothenburg.

Konturek, P., Konturek, S. and Hahn, E. 2004. Duodenal alkaline secretion: its mechanisms and role in mucosal protection against gastric acid. Dig. Liver. Dis. 36(8), 505–512; doi:10.1016/j.dld.2004.03.008

Krauss, R.S., Chihara, D. and Romer, A.I. 2016. Embracing change: striated-for-smooth muscle replacement in esophagus development. Skeletal. Muscle. 6(1), 27; doi:10.1186/s13395-016-0099-1

Lin, C., Liu, W., Xie, J., Li, W. and Zhou, Z. 2020. The lubricating function of mucin at the gastroscope device–esophagus interface. Tribol. Lett. 68(3), 82; doi:10.1007/s11249-020-01322-9

Liu, Y., Li, D., Ma, L., Wen, Y. and Shi, D. 2025. The barrier and protective functions of intestinal mucin in defense against Candida albicans. Front. Microbiol. 16, 1561004; doi:10.3389/fmicb.2025.1561004

Livak, K.J. and Schmittgen, T.D. 2001. Analysis of relative gene expression data using real-time quantitative PCR and the 2−ΔΔCT method. Methods 25(4), 402–408; doi:10.1006/meth.2001.1262

Luu, Y., Junker, W., Rachagani, S., Das, S., Batra, S.K., Heinrikson, R.L., Shekels, L.L. and Ho, S.B. 2010. Human intestinal MUC17 mucin augments intestinal cell restitution and enhances healing of experimental colitis. Int. J. Biochem. Cell Boil. 42(6), 996–1006l; doi: 10.1016/j.biocel.2010.03.001

Mackie, R.I. 2002. Mutualistic fermentative digestion in the gastrointestinal tract: diversity and evolution. Integr. Comp. Biol. 42(2), 319–326; doi:10.1093/icb/42.2.319

Nakatsuji, S., Szabo, K.A. and Elmore, S.A. 2018. Small and large intestine. In Boorman’s pathology of the rat. Amsterdam, The Netherlands: Elsevier / Academic Press, pp: 51–69; doi: 10.1016/B978-0-12-391448-4.00007-1

.Nzalak, J.O., Onyeanusi, B., Samuel, A.O., Voh, A.A. and Ibe, C. 2010. Gross anatomical, histological and histochemical studies of the esophagus of the African giant rat (Cricetomys gambianus). J. Vet. Anatomy 3(2), 55–64; doi:10.21608/jva.2010.44900

Sary, S., Dougbag, A., Zaghloul, D., Mohamed, K. and Gomaa, M. 2025. The histological structure of chief cell of fundic gland region of one-humped camel (Camelus dromedarius). Alex. J. Vet. Sci. 84, 1–7; doi:10.5455/ajvs.208249

Schmidt-Nielsen, K. 1997. Animal physiology: adaptation and environment. Cambridge, United Kingdom: Cambridge Univ Press.

Slomiany, B. and Slomiany, A. 1991. Role of mucus in gastric mucosal protection. J. Physiol. Pharmacol. 42(2), 147–161.

Treuting, P.M., Dintzis, S.M. and Montine, K.S. 2017. Comparative anatomy and histology: a mouse, rat, and human atlas. Amsterdam, The Netherlands: Academic Press.

Uehara, T., Elmore, S.A. and Szabo, K.A. 2018. Esophagus and stomach. In Boorman’s pathology of the rat. Amsterdam, The Netherlands: Elsevier / Academic Press, pp: 35–50; doi: 10.1016/B978-0-12-391448-4.00006-X

Wattel, W. 1978. Structural and functional development of rat and mouse gastric mucous cells in relation to their proliferative activity. Rijksuniversiteit Utrecht.

Withers, P.C., Cooper, C.E., Maloney, S.K., Bozinovic, F. and Cruz-Neto A.P. 2016. Ecological and environmental physiology of mammals. New York, NY: Oxford Univ Press; doi:10.1093/acprof:oso/9780199642717.001.0001

Yandrapu, H. and Sarosiek, J. 2015. Protective factors of the gastric and duodenal mucosa: an overview. Curr. Gastroenterol. Rep. 17(6), 24; doi:10.1007/s11894-015-0452-2