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Ex vivo ultrasonographic and histological morphometry of small intestinal wall layers in horses

Abstract Ultrasonographic morphometry of wall layers is commonly used in veterinary patients with suspected small intestinal disease, however published studies comparing this method with histopathology in horses are limited. This prospective, methods comparison study compared the qualitative and quantitative characteristics of small intestinal wall layers using ex vivo high‐frequency ultrasound versus histopathology in a sample of 16 horses. Transverse section images of duodenum, distal jejunum, and ileum were acquired with a high‐frequency linear transducer (7–15 MHz). Transverse histological cryosections were obtained at the same level. Appearance and measurements of the intestinal wall layers were assessed on the ultrasonographic and histological images. High‐frequency scanning with the probe in close contact with the serosal surface of the equine intestinal wall allowed a clear and detailed definition of wall layers. A hyperechoic line was consistently detected within the tunica muscularis in all the intestinal tracts, corresponding histologically to the interface between its longitudinal and circular muscle layers. The overall trend of the values for wall layers thickness was comparable between ex vivo ultrasonography and histology. However, a poor agreement was found between the two methods for all layers. The ultrasonographic measurements were thicker compared to histological measurements, with the exception of the total wall and the muscular layer thicknesses. These layers were thinner on ultrasonography in the duodenum and in all the intestinal segments, respectively. Findings from the current study can be used as background for future ultrasonographic investigations of small intestinal diseases in horses.


INTRODUCTION
Ultrasonography (US) is one the most commonly used non-invasive diagnostic tools in equine practice 1,2 and can be particularly useful for the diagnosis of gastrointestinal disease. In fact, this technique allows the evaluation of intestinal diameter, wall thickness and layering, luminal contents, and motility. 2 These parameters can be altered during intestinal diseases. 1,3 Intestinal wall thickness may increase in the setting of different inflammatory/infiltrative bowel diseases, strangulating intestinal lesions, or edema. 1,2,4 On the other hand, the pathological thinning of the intestinal wall has not received specific consideration in horses so far. The normal US appearance and reference ranges of the overall intestinal wall thickness in the different GI tracts have been widely reported in horses. 1,5 Based on previous publications in small animals, 6,7 the equine ultrasonographic intestinal layers were assumed to correlate well with the histological layers (mucosa, submucosa, muscularis, and serosa), but few research have been performed on ex vivo equine intestine. [8][9][10] More recently, only one study describes the ultrasonographic thickness of each small intestinal wall layer of the equine and its good correlation with histological measurements of any segment, except for the ileum. 8  This study aimed to assess wall layering and thickness of the different small intestinal segments by ex vivo high-resolution US, and to evaluate the agreement between ultrasonographic and histologic measurements of the individual layers and overall small intestinal wall in a sample of healthy horses. We hypothesized that wall thickness measurements obtained by this method at the different intestinal levels would agree with wall layering morphometry determined by histology as a reference method.

Experimental design and sample collection procedures
The study was a prospective, methods comparison design. Twentythree horses (14 males and 9 females), of various breeds, aged 1.5-24 years were initially recruited from a local abattoir (Emilia Romagna region, Italy) during the period of March-April 2019. No ethical approval was required as all the horses were presented to the abattoir for reasons unrelated to the study. Horses were deemed free of gastrointestinal disease and included in the study on the basis of the veterinary inspection report at the abattoir, normal CBC and serum biochemistry, and absence of gross, ultrasonographic, and histological lesions at the different sections of the small intestine. Collected samples that did not fulfill all the inclusion criteria were excluded from the analysis.
Blood and tissue collections were performed by the veterinary personnel responsible for animals and meat inspection at the abattoir and by an equine internal medicine specialist (FF). Jugular blood samples were taken at the time of exsanguination and kept refrigerated till analysis (within 2 h). A complete blood count (CBC; Advia 2120, Siemens Healthcare Diagnostics, Tarrytown NY, USA) and routine serum biochemistry (U400, Olympus/BeckmanCoulter, Brea CA, USA) were performed. Two segments of the gastrointestinal tract were resected from each animal immediately (no longer than 30 min) after slaughtering.
The first intestinal segment included the pyloric portion of the stomach to the junction between the duodenum and jejunum and the second the caudal portion of the jejunum to the ileocecal junction (ICJ), including the base of the cecum. The small intestinal samples consisted of 25 cm in length segments harvested from the descending duodenum (40 cm far from the pyloric sphincter), caudal jejunum (150 cm far from the ICJ), and ileum (20 cm far from the ICJ). The samples were immediately wrapped in moist towels and plastic for transport, hence ultrasound exam and processing were performed within two hours. The samples were coded and randomized for blinded ultrasonographic and histological evaluations. All ultrasound examinations were conducted by the same sonographer (AD), a veterinarian with more than 15 years of experience using a real-time ultrasound machine (Philips © IU22, Ultrasound System, Philips Healthcare, Monza, Italy) equipped with a high-frequency

Evaluation and measurements of ultrasonographic images
The DICOM images were transferred to a computer workstation (Imac, macOS Monterey version 12.0.1, Cupertino CA). Open-source image analysis software (OsiriX Imaging, Trillium Technology, Ann Arbor, MI) was used for subsequent image analyses. The images were displayed on a calibrated, 2-megapixel high-brightness monitor (Eizo RX240; Cusano). All images were assessed by the same investigator who had previously acquired the sonographic images (AD), without knowledge of histological measurements. In all images, the overall ultrasonographic wall layering was subjectively assessed along with measurements of the total wall and layer thicknesses.
Electronic calipers were placed at the outside edge of each individual intestinal layer and leading edge-to-leading edge measurements were made from the serosal interface to the luminal interface of the mucosa ( Figure 1C). Three consecutive measurements of mucosal, submucosal, muscularis, serosal, and total intestinal wall thickness between the intestinal folds were obtained from the images of PBSfilled duodenum, jejunum, and ileum. Repeated measurements of the individual layers and of the total intestinal wall were subsequently averaged.

Statistical analysis
Only samples matching all inclusion criteria underwent the subsequent analysis, and measurements by the two methods were paired.

RESULTS
The study included 16 ( jejunum, and 15 ileum) were included in the analysis.

Subjective evaluation of ultrasonographic and histological images
Ultrasonographic evaluation showed a typical layering of the intestinal wall, in which five layers were easily identified from outside to inside in both the empty and distended intestine: serosa (hyperechoic), muscularis (hypoechoic), submucosa (hyperechoic), mucosa (hypoechoic), and mucosal surface (hyperechoic) (Figure 1). Furthermore, all intestinal samples showed an additional hyperechoic thin line parallel to the submucosa and serosa, in the outer aspect of the ultrasonographic muscular layer ( Figure 3A). On histology, no morphological abnormalities and the characteristic layered stratigraphy were found in all the samples.

Ultrasonographic and histological measurements and agreement
Results of the ultrasonographic and histological measurements of the total wall thickness and individual wall layers in the different intestinal segments as well as of comparisons of the US measurements between the different segments are reported in Table 1. The relative contribution (%) of each individual wall layer to the total wall thickness was consistent between US and histology and among the three intestinal segments, and was represented in such decreasing order: muscularis, mucosa, submucosa, and serosa (Table 2).
A significant difference was found in the ultrasonographic measurement of the total intestinal wall and that of the serosa, submucosal, and muscular layer in the three different segments (overall P < 0.001, P = 0.019, P = 0.003, and P < 0.001, respectively), whereas no difference was found for the mucosa (P = 0.305; Figure 5). In particular, the  total wall thickness of the ileum was higher compared to that of the duodenum and ileum (P < 0.01 for both comparisons). Furthermore, the serosal and submucosal layers of the ileum were higher compared to that of the duodenum (P < 0.05 and P < 0.01, respectively), and the ileum muscular layer was higher compared to that of the duodenum and jejunum (P < 0.001 and P < 0.01, respectively).

US (µm) Histo (µm) US (µm) Histo (µm) US (µm) Histo (µm)
The agreement between ultrasonography and histology was poor for all the measurements of both the total intestinal wall and each layer thickness in the three intestinal segments (Supporting Information 1).
In particular, the ultrasonographic measurements were all thicker compared to the histological ones, with the exception of the total wall and

DISCUSSION
The intention of the current study was to compare ex vivo ultrasonographic and histologic characteristics of small intestine layers in a sample of clinically normal horses. The ex vivo ultrasonographic appearance of the five wall layers (serosa, muscularis, submucosa, mucosa, and mucosal surface) of the equine small intestine in the current study was consistent with findings from previously published descriptions. [3][4]9,12 In those previous studies, transducer frequencies ranging from 2.5 MHz to 7.5 MHz were employed. The use of a highfrequency transducer allowed to achieve a higher resolution of the wall layering. In particular, a further thin hyperechoic line within the muscular layer, representing the interface between the inner and outer parts of the tunica muscularis, was observed along all the equine small intestine as also described in humans [13][14][15] and dogs. 5  With regard to US measurements, in the present study, the small intestinal wall thickness was at the higher limit or slightly over compared to previous reports using low to medium frequency probes. 1,[3][4][5]10,11,25,26 Transabdominal and transrectal US in horses encounter many anatomical and technical limitations to intestinal walls' imaging quality (e.g., size of the abdomen, distance of the viscera from the contact surface, low-frequency probes). A thinner small intestinal wall compared to the present findings has been reported scanning ex vivo intestine using a 7.5 MHz probe, 9 and also a high fre-quency (10-18 MHz) probe. 8  Comparing US measurements of the different intestinal segments, the overall wall thickness of the ileum resulted thicker than the more proximal ones. There are no in vivo US reference ranges specifically referred to the ileum for direct comparison, likely reflecting that it is limitedly accessible for measurement in vivo. However, our finding was in line with a previous ex vivo study 9 and with the recent study by Bevevino et al. 8 Along with the above-mentioned technical factors, sampling at the level of the distal jejunum versus more proximal jejunal sites is most likely key in justifying the relatively higher wall thickness of the jejunum compared to duodenum recorded in the current study but not by Bevevino et al. 8 This is mainly reflected by a thicker muscular layer present at a distal level, gradually nearing to resemble the wall stratigraphy of the ileum. This is particularly evident in histology (Supporting Information 3), and also renders reason for the less usual US appearance of the jejunum showed in the current study (Figure 1). While the clinical relevance of this difference is undefined, we should be aware that such differences in jejunal wall morphometry and appearance could be encountered during a US assessment of the small intestine, especially in optimal conditions. Thickness ranges for the individual wall layers were recently provided at the level of duodenum, jejunum, and ileum by high-frequency US. 8 In the present study, among the different segments, the US muscular layer of the ileum was significantly thicker than in the other tracts; despite a distinction being visible, the two muscular layers could not be separately measured with accuracy.
The same finding was reported only at the level of the distal ileum in dogs and cats. 7,29 Differently, Worth 10 did not report a prominence of this layer with respect to the others in the distended viscus. Consistently, the tunica muscularis was the most developed in the ileum by histology. This is consistent with the functional specialization of this trait in overcoming the resistance of the ileocecal valve and generating vigorous peristaltic movements. 30 In their relative contribution to the ultrasonographic wall thickness, the muscular layer was followed by the mucosal, then the submucosal, and last the serosal one, and this pattern was consistent along the different intestinal segments. On the contrary, differences in the thickness of individual layers between segments have been reported in small animals by US. 7,29,31,32 Submucosal layer of the ileum was higher compared to that of the duodenum. The findings were mainly attributed to a major contribution of villi and Brunner's glands, and of Peyer's patches, in duodenum and ileum, respectively. Such structures were similarly represented in our samples, albeit did not affect the relative thickness between tracts. Nevertheless, in histological samples from the younger horses, the large Peyer's patches in the ileal mucosa and submucosa were occasionally themselves.
Despite the overall trend of the variables being consistent between the two methods, there was a poor agreement between US and histologic measurements for the total wall and individual layers in all the segments. In contrast to our results, a good agreement between ex vivo US and histologic measurements of the wall thickness and different layers of the small and large intestinal segments, except for the ileum, has been reported in horses. 8 However, differences in the setting and methods, both operative and analytic (e.g., statistical setting) between the two studies have to be considered for the discrepancy.
Overall, although we were not able to statistically support our hypothesis, the clinical relevance of the discrepancy is uncertain as morphology and trends were consistent between methods. Nevertheless, sim-ilar to our findings, the ultrasonographic intestinal layers have been reported to not directly correspond to the histological intestinal layers in human literature. 13,33 The lack of correlation has been predominately attributed to three factors. First, during histological processing, tissue may lose its original size, which may be somehow layer dependent. Fixation and paraffin embedding cause a considerable retraction/shrinkage of the intestine. 34 In the present study, we were not able to avoid tissue retraction due to the fixation, but cryosectioning spared the tissues from further shrinkage due to paraffin embedding.
To note, the larger difference in measurements between US and histology occurred for the mucosa in all segments. In a similar study in cats, it has been previously hypothesized that, due to its composition, tissue shrinkage and protein coagulation due to alcohol-based fixatives could be more pronounced at the mucosal level. 6 However, we employed a non-alcohol-based fixative, so it is unknown whether this could be similarly a critical factor in our study. As a second factor, echogenic bands are generated by an interface between structures of different acoustic impedance (e.g., gastrointestinal wall layers) and should not be considered to necessarily correspond to an anatomical layer. 33 Moreover, the ultrasonographic axial resolution, which is mainly limited by the spatial pulse length (SPL) of the instrument, should be considered. As a result, even though the interface between two intestinal layers may have no histological thickness, the hyperechoic interface produced between them will be at least the same thickness as the SPL. 13,33,35 Interestingly, all ultrasonographic measurements were thicker than the histological ones in all the intestinal segments, except for the total wall of the duodenum and the muscular layer of all segments, where the opposite occurred. The same finding was reported for the muscular layer at the level of the distal ileum in cats, being only this layer thicker by histology than by US. 6 Martinez et al 6 hypothesized that the thickness of the muscular layer on US is generated by the thickness of the tunica muscularis minus the interface between the submucosa and the tunica muscularis (which would be added to the submucosa instead), explaining a thinner muscularis layer by US than by histology. Thus, this process would play in favor in the case of the mucosa, submucosa, and likely the serosa, with their relative external and internal interfaces, potentially contributing to the greater ultrasonographic than histologic thickness of these layers, and to their poor agreement.
On the other hand, taking measurements on enlarged microphotograph of appropriately labeled tissue sections allowed the accurate defining of the thickness of all individual layers. For instance, Neu-roTrace labeling permitted an accurate definition of the thickness of the mucosa and in particular the outer limit of the muscularis mucosae.
Another issue that might be considered is the impossibility to reproduce the same section plane by US and histology. Furthermore, with regard to the mucosa, it is very difficult to constantly obtain a histologic section parallel to the villi, which will represent the full height of this layer. Last, while a circular section of the organ is measured by US, the section is plane on histology, employing a degree of stretching and flattening of the surface.
The study has several limitations. The study setting did not allow collecting a complete history and clinical picture of the horses; however, in the authors' opinion the US, gross and histologic assessment of the small intestine were adequate to the study purpose. The overall sample size was limited and could not be stratified by age, body weight, breed, or management in relation to the US and histological morphometric findings. In the authors' opinion, all subjects may be considered as presenting a fully developed digestive system. This might be worth of future insights in view of a diagnostic or prognostic application. Despite our effort to minimize the time between samples collection and processing, at least 90 min were needed to complete the procedure, and some lag occurred between the first and the last sample, which could potentially contribute to variability in wall thickness. No post-mortem effect on equine small intestinal length within a few hours has been reported. 36 However, dehydration of the mucosa, which induced loss of turgidity, stiffness, and a thinning of intestinal villi, was reported after 40 min in rats. 37 In conclusion, the findings of the current study partially supported those reported in a recent publication on a similar topic. 8 Highfrequency US with the probe in direct contact with the intestinal wall provided detailed visualization of wall layering that was comparable to qualitative histological features. However, in our study, the agreement between methods for most quantitative measures of wall thickness was not statistically significant, likely due to histological preparation factors such as the degree of bowel distension and sample processing.

ACKNOWLEDGEMENT
Open Access Funding provided by Universita degli Studi di Bologna.