Achalasia is a disease of the esophagus, characterized by impaired relaxation of the lower esophageal sphincter (LES) and absent peristalsis in the smooth muscle esophageal body, resulting in symptoms of dysphagia, regurgitation, chest pain and weight loss.¹ It affects approximately 20/100 000 persons in the North American population.²

Achalasia is separated into 3 distinct subtypes. While all subtypes involve impaired LES relaxation, type 1 is associated with minimal contractility in the esophageal body, while type 2 is associated with episodes of pan-esophageal pressurization and type 3 with spastic esophageal contractions.³

The etiopathogenesis of achalasia is poorly understood, partly owing to difficulty obtaining specimens of esophageal smooth muscle from patients with the disease, as well as a lack of suitable animal models. Pathologically, there is a loss of inhibitory nitric oxide (NO) synthase-producing neurons in the myenteric plexus associated with a marked lymphocytic infiltration of the nerve plexus and fibrosis.⁴ However, little is known about the intrinsic innervation of smooth muscle in this disease nor the changes that occur in the muscle following loss of innervation. The histopathology is complicated by the existence of 3 subtypes of achalasia that appear to vary with regards to neuron loss, fibrosis, and decrease in number of interstitial cells of Cajal.^5,6 It is also unclear if these differences are true when comparing the esophageal body to the LES.

In recent years peroral endoscopic myotomy (POEM) has proven to be a safe and effective minimally invasive therapy for all forms of achalasia.⁷ This procedure involves a tunneled dissection of the submucosa into the gastric cardia whereby a circular smooth muscle (CSM) myotomy is completed. POEM provides the endoscopist the ability to directly biopsy the CSM layer of the esophagus, thereby allowing access to a tissue compartment that previously required invasive surgery. Using biopsies from patients undergoing POEM, we sought to characterize the CSM, using advanced immunocytochemical techniques to evaluate the neuronal and smooth muscle populations. In light of the established correlation between inflammation-driven hyperplasia of intestinal smooth muscle and epigenetic modulation of phenotype,⁸ we were also interested in characterizing esophageal CSM phenotype and innervation according to location in the esophageal body, and further, to explore, in a preliminary fashion, possible differences among the subtypes of achalasia.

Study Design

This study was conducted as an observational case control analysis comparing patients with achalasia undergoing a POEM procedure with individuals without achalasia who underwent partial esophagectomy for management of proximal gastric cancer where the esophagus was not involved. Consecutive achalasia patients were enrolled between June 2017 and February 2019, with a further 5 patients enrolled in December 2020 for a separate part of the study.

Patients were included if they were > 18 years old and had a confirmed diagnosis of achalasia with high-resolution esophageal manometry and were deemed medically suitable to undergo POEM.

Retrieval of Human Esophageal Tissue Samples

During the POEM procedure standard endoscopic biopsy forceps were used to obtain 2 samples of CSM (identified by the direction in which the muscle fibers are oriented during POEM) from the proximal margin of the myotomy (PE) and 2 from the LES. They were then placed directly into neutral buffered formalin and embedded in paraffin according to established procedures.⁹ Similarly, samples from 5 other patients were separately placed in RNAlater stabilization solution (Fisher Scientific, Nepean, ON, Canada) for subsequent retrieval of RNA. Control tissues were obtained from patients > 18 years of age with no known esophageal disease during distal esophagectomy for proximal gastric cancers and adjacent healthy tissue prepared in an identical fashion.

Immunocytochemistry

To assess for tissue morphology, paraffin-embedded control and POEM specimens were sectioned and stained with H&E according to previously established protocols.¹⁰

To detect axon structure, antibodies to pan-neuronal marker protein gene product (PGP) 9.5 and the nitrergic phenotypic marker neuronal nitric oxide synthase (nNOS) were used (both 1:200; Cell Signaling Technology, Inc, Beverly, MA, USA). Antibodies to transgelin (SM22) (1:2000; Abcam, Waltham, MA, USA), α-smooth muscle actin (SMA) (1:1000; MilliporeSigma, Burlington, ON, Canada), and smoothelin (SMOO) (1:2000; Abcam) were used to label smooth muscle cells. Anti-Ki67 antibodies were used to test for the presence of proliferating nuclei (1:100; Bio-Techne, Etobicoke, ON, Canada). Mast cells were detected with anti-mast cell tryptase (1:100; Agilent Dako, Santa Clara, CA, USA). All antibodies were diluted in phosphate buffered saline with 0.2% Tween-20 (PBS-T), and where necessary, tissue was incubated with citrate buffer (pH 6) at 95°C for 20 minutes or with 1× target retrieval solution (Dako) according to the manufacturer’s directions. Slides were then incubated with 1% goat serum in PBS-T for 1 hour to block nonspecific binding before overnight incubation in primary antibodies at 4°C, followed by incubation in the appropriate Alexa-linked secondary antibodies (MilliporeSigma) for 2 hours at room temperature. Cell nuclei were identified with Hoechst (1 µg/mL; MilliporeSigma), then staining was visualized with fluorescence microscopy (BX51; Olympus, Shinjuku, Tokyo, Japan). ImageJ (version 1.53; NIH, Bethesda, MD, USA) was used for smooth muscle marker staining intensity analysis, and Image Pro Plus version 6 (Media Cybernetics, Inc, Rockville, MD, USA) was used for axon area, axon intersections per CSM nuclei, and mast cell quantification. A minimum area of 5 µm² was used for inclusion in axon measurements. At least 3 non-adjacent sections of the CSM layer in control tissue or biopsy tissue were analyzed under standardized exposure settings, averaged, and normalized to an on-slide control sample to ensure accurate and reproducible densitometry and area analysis.

Sirius Red Staining

Adjacent sections to those used for immunocytochemistry were treated with Sirius Red/Fast Green (BDH Chemicals) differential stain to allow analysis of the proportion of collagen to cellular protein as described.⁹ At least 3 non-adjacent images of each stained sample were acquired by light microscopy (Olympus BX60; Infinity 2 Camera; Infinity Capture [Lumenera Corporation, Ottawa, Canada]). These were used to measure the proportions of collagen (red) and non-collagen (green) protein (Image Pro Plus version 6; Media Cybernetics).

Quantitative Real-time Polymerase Chain Reaction

RNA was extracted from esophageal tissue using the E.Z.N.A Total RNA Kit I (Omega Bio-Tek, Inc, Norcross, GA, USA) according to the manufacturer’s instructions. An iScript cDNA Synthesis Kit (Bio-Rad Laboratories, Ltd, Mississauga, ON, Canada) was used to convert RNA to cDNA, and the complete reaction mix was incubated in a thermal cycler: priming for 5 minutes at 25°C, reverse transcription for 20 minutes at 46°C, and RT inactivation for 1 minute at 95°C. Quantitative polymerase chain reaction (qPCR) was used to measure changes in SMA, SM22, SMOO, myosin heavy chain 11 (MYH), type I collagen (COL1), type III collagen (COL3), and glial cell line-derived neurotrophic factor (GDNF) mRNA expression in human esophageal control and achalasia samples (primer sequences; Supplementary Table 1). These values were normalized to β-actin mRNA levels, after its PCR amplification efficiency was confirmed. Reactions were carried out in a StepOnePlus Real-Time PCR system (Applied Biosystems, Inc, Woburn, MA, USA), held for 10 minutes at 95°C, and cycled for 15 seconds at 95°C and 60 seconds at 60°C for 40 cycles. Relative changes in mRNA expression were determined by comparative threshold (Ct) analysis using the ΔΔCt method of relative quantification and are presented as the average independent outcomes.

Statistical Methods

Quantitative data are expressed as means ± standard deviation. Differences were considered significant for P-values < 0.05 using ANOVA or t test where appropriate. Spearman correlation was performed to determine whether certain continuous variables correlated with immunocytochemical values.

Ethics

All patients were provided a consent form describing the study in detail and with sufficient information to make an informed decision about their participation. The consent was signed by the patient or their legally authorized representative. The study was approved by the Queen’s University Research Ethics Board (#6020138).

Patient Demographics

Demographics of the initial 25 enrolled patients are summarized in Table. Average patient age was 50.4 years, and roughly half of the patients were male. Most patients had type 2 achalasia (18), followed by type 1 (4) and type 3 (3). Supplementary Table 2 summarizes the characteristics of the 5 additional achalasia patients whose biopsies were obtained for PCR analysis.

Table. Patient Demographics

Patient factor
Sample size	25
Age (yr)	50.4 ± 18.1
Sex (male)	13
Subtype of achalasia Type 1 Type 2 Type 3	4 18 3
Duration of symptoms (yr)	7.8 ± 10.4
Eckardt score	7.1 ± 2.3
Previous invasive treatments (no. of patients) None Botox Balloon/Savary dilation Pneumatic dilation Heller myotomy Multiple above therapies	16 2 3 2 3 5

Mean values are presented as n or mean ± SD.

General Histology of Biopsy Tissues

Hematoxylin and eosin staining of paraffin sections of pinch biopsies targeting the CSM confirmed their smooth muscle nature. A comparison of tissue from control patients (n = 3) and those with achalasia showed specimens were similar in appearance with no significant electrocautery artifact present in any samples (Fig. 1A). There were no prominent differences in inflammatory infiltrate, with the number of polymorphonuclear leukocytes or eosinophils in the CSM layer being similar. Nonetheless, more areas of CSM tissue appeared non-cellular (ie, composed of extracellular matrix) in the biopsy specimens from achalasia patients than in controls (eg, Fig. 1A), and application of further, more detailed analyses (see below) showed significant and characteristic distinctions.

Figure 1. Histopathology of peroral endoscopic myotomy biopsies of esophageal circular smooth muscle (CSM) showing diminished innervation in achalasia. (A) Representative images of CSM from the lower esophageal sphincter stained with H&E from control tissue and achalasia. Scale bar, 20 µm. (B, C) Detection of innervation of CSM cells by immunocytochemistry, with representative images of staining with the pan-axonal marker protein gene product (PGP) 9.5 (B; red staining) in control and achalasia tissue. (C) Images showing the subset of nitrergic axons in the CSM by staining with antibodies to neuronal nitric oxide synthase (nNOS, green). Scale bar, 20 µm. Arrows and arrowheads indicate positive staining. Nuclei are stained with Hoechst (blue). (D) Image analysis showing decreased percent total area of positive staining per image for PGP or nNOS in achalasia (Ach) vs control (Con). *P < 0.0001 vs control. (E) Graph depicting the percent area of positive staining for PGP in each subtype of achalasia.

Innervation of Circular Smooth Muscle Is Deficient in Achalasia

As expected, analysis of the CSM layer of tissue from control or achalasia biopsy tissues showed no PGP 9.5-labeled neuronal cell bodies in any sample, although cell bodies of the myenteric plexus were detected below the CSM in the sections of full-thickness control tissues (not shown). In control tissues, the use of anti-PGP antibodies showed abundant labeling of axons in the CSM layer (Fig. 1B).

Image analysis was used to quantify axon number as well as smooth muscle cell number and image area in each section, revealing that control tissues averaged 3.5 ± 0.1 axons/10 CSM, or 0.8 ± 0.3% of the total CSM area (n = 3; Fig. 1D). As predicted, detection of the nitrergic subtype of axons using anti-nNOS antibodies showed fewer axons, which averaged 1.1 ± 0.1 axons/10 CSM, or 0.2 ± 0.0% of the total CSM area. Therefore, the nitrergic axons, representing inhibitory innervation, represented 29.8 ± 1.6 % of the total axon number.

Comparison of these characteristics between control and achalasia tissues showed that the number of axon processes in achalasia samples was substantially reduced compared to control, with many biopsies entirely devoid of PGP labeling (Fig. 1B and 1C). Overall, there was an average of 0.16 ± 0.03 axons/10 CSM in the LES region and 0.23 ± 0.09 axons/10 CSM in the PE. No PGP positive axons were detected in 1 biopsy from type 1 or 4 biopsies from type 2, despite analysis of several non-adjacent sections, and values were independent of subtype (Fig. 1E; P > 0.05). In all cases, labeling was confirmed using on-slide control sections.

As expected, nitrergic axons were virtually absent from biopsies of the achalasic esophagus, with nNOS labeling detected in only 5 of the 25 biopsies from the LES region and 4 of the 25 biopsies from the proximal region. This ranged from 0.04 to 0.16 axons/10 CSM. Similarly, positive antibody labeling was visualized on on-slide control sections to verify successful technical outcomes. This analysis confirmed the significant decrease or loss of innervation in the CSM region of both the LES and the PE in achalasia.

Mast Cell Numbers Are Increased in Achalasia Biopsies

An increased presence of mast cells is characteristic of previous or ongoing inflammation, and an increase within achalasia biopsies has been reported.^11,12 In contrast to previous studies, we quantified these cells as number/area of CSM analyzed, which thus allowed direct comparison among groups. In the control CSM layer, few mast cells were detected, averaging 1.0 ± 0.3 cells/10⁴ µm² (Fig. 2; n = 3). In contrast, more mast cells were detectable in achalasia, with this being similar between the LES and the PE tissues (Fig. 2; 3.1 ± 0.6 and 3.7 ± 0.6 cells/10⁴ µm² for LES and PE, respectively). There was variation among the samples, with no mast cells detected in the LES of 3 patient samples, and more than 10 cells/10⁴ µm² in 2 of the samples. There did not appear to be a correlation with mast cell presence in the subtypes of achalasia or with respect to patient characteristics, such as age, duration of symptoms and Eckhardt score (not shown).

Figure 2. Biopsies from achalasia show a characteristic presence of mast cells in the circular smooth muscle (CSM). (A, B) Images of immunocytochemistry for anti-mast cell tryptase (green) showing mast cells (eg, arrows) in the CSM from control (A) or achalasia tissue (B). Nuclei, blue. Scale bar, 20 µm. (C) Increased mast cell number per 10⁴ µm² in achalasia vs control CSM. *P < 0.05 vs control.

Smooth Muscle Phenotype Is Altered in Achalasia

In animal models, inflammation that causes damage and loss of innervation of intestinal smooth muscle is correlated with the onset of hyperplasia of the target smooth muscle cells; proliferation ends when innervation is restored.^13,14 To test whether the denervated smooth muscle cells in achalasia were actively proliferating at the time of POEM, we labeled tissue with antibodies to Ki67, an established marker of ongoing cell division in human tissue.¹⁵ While control mucosa showed prominent nuclear labeling of Ki67 in dividing epithelial cells, the CSM region showed no expression of this marker, despite analysis of several non-adjacent sections for each tissue (Fig. 3D). Similarly, analysis of non-adjacent sections of biopsy tissue from achalasia patients showed complete absence of Ki67 expression in all tissues (n = 25), showing that the denervated esophageal CSM cells were not actively proliferating at the time of biopsy.

Figure 3. Smooth muscle phenotype is altered in achalasia, with no evidence of ongoing smooth muscle proliferation. (A-C) Representative images of esophageal circular smooth muscle (CSM) from control and achalasia showing antibody staining for the smooth muscle marker proteins, transgelin (SM22) (A), α-smooth muscle actin (SMA) (B), and smoothelin (SMOO) (C) expression. Scale bar, 20 µm. (D) Antibody staining for the proliferation marker Ki-67 showing staining in positive control tissue (esophageal mucosa) but absence from control and achalasia esophageal CSM. Scale bar, 20 µm. (E) Image analysis showing staining intensity of SMA, SM22 and SMOO in CSM biopsies from achalasia patients (Ach) relative to control (Con). *P < 0.05 vs control. (F) Staining intensities of SMA relative to control in the 3 subtypes of achalasia. *P < 0.05 vs control.

To assess smooth muscle phenotype directly, we labeled sections of control tissue with antibodies to the smooth muscle markers SMA, SM22 and SMOO.^16,17 There was no significant difference among the control tissues in terms of intensity of staining, nor of intensity in the circular vs longitudinal smooth muscle compartments (data not shown). Furthermore, there was no difference between the PE or LES in terms of intensity of fluorescence. Therefore, we averaged the image intensity of these 2 sites and compared this with the staining intensity in achalasia samples. Interestingly, there was a significant increase in expression levels of both SMA and SMOO but not SM22 in the CSM of achalasia biopsies as compared to control (Fig. 3). Analysis showed that the relative intensity of labelling was 1.39 ± 0.27 vs 1.00 ± 0.04, P < 0.05 and 1.32 ± 0.10 vs 1.00 ± 0.1 for SMA and SMOO, respectively (Fig. 3E). We then compared staining intensity among the 3 types of achalasia and noted a significant increase in SMA but not SM22 or SMOO labelling intensity in Type 1 as compared to Type 2 and Type 3 (SMA: 1.82 ± 0.45 vs 1.33 ± 0.18 vs 1.18 ± 0.15, P < 0.05; Fig. 3F). This suggests that the esophagus of achalasia patients may show aberrant contractile smooth muscle marker expression, which may contribute to abnormal function.

To test whether these phenotypic changes correlated with patient factors, we performed a correlation analysis. This showed that the increase in SMA and SMOO staining in the achalasia biopsies correlated with duration of symptoms (Fig. 4), although not with Eckhardt score nor age, (after identifying and removing a single outlier by Grubb’s test). Interestingly, this outlier had numerous procedures to their esophagus, including repeated Heller myotomies and pneumatic dilations raising the possibility that repeated instrumentation can modify muscle physiology.

Figure 4. Positive correlation of the staining intensity of the smooth muscle markers, α-smooth muscle actin (SMA) and transgelin (SM22) with duration of symptoms. Data depict the intensity of SMA and SM22 staining per patient biopsy correlated with the duration of reported patient symptoms. For SMA, Pearson r = 0.50; 95% CI, 0.12 to 0.75; R² = 0.26, P < 0.01. For SM22, Pearson r = 0.42; 95% CI, 0.01 to 0.70; P < 0.05.

To detect a genetic basis for the altered smooth muscle cell phenotype, a subset of biopsy samples from 5 additional achalasia patients (1 type 1 and 4 type 3) was analyzed by qPCR to detect differences in expression of mRNA for SMC markers (Fig. 5). For this, primers for SMA, SM22, SMOO, and MYH were used to amplify mRNA levels relative to mRNA for β-actin in samples from achalasia vs control. While MYH mRNA expression showed a decrease relative to control, levels of SMA, SM22 and SMOO generally increased relative to controls. For example, SM22 levels ranged from 1.2× to 2.7× control, while SMOO mRNA levels ranged from 4.0× to 21.5× above control levels (P < 0.05). In addition, we tested for the presence of the neurotrophic factor GDNF, essential for myenteric neuron survival and axonal innervation of SMC.^18,19 We found increased expression of GDNF mRNA in biopsies relative to control (P < 0.05, Fig. 5).

Figure 5. Outcomes of quantitative polymerase chain reaction for expression of mRNA in circular smooth muscle (CSM) for smooth muscle marker genes (α-smooth muscle actin [SMA], transgelin [SM22], smoothelin [SMOO], and myosin heavy chain 11 [MYH]), type I collagen (COL1) and type III collagen (COL3) and the enteric neurotrophin GDNF, expressed per patient vs control. *P < 0.05 increased relative to control; ^#P < 0.05 decreased relative to control.

While previous studies have reported on tissue fibrosis in samples from patients with achalasia,¹² there are no statistical comparisons with control tissues. In the current study, we measured mRNA levels of COL1 and COL3 mRNA levels in biopsy samples (n = 5) relative to control (n = 3) (Fig. 5). The mRNA for COL3 was reduced in the biopsy tissue (P < 0.05) while the levels of COL1 were increased relative to control but did not reach statistical significance.

Next, we examined evidence for fibrosis in achalasia, using histochemistry for collagen (Sirius Red staining) in control and biopsy samples to detect the relative amounts of collagen (Fig. 6). Representative images from control, type 1 and type 3 achalasia are shown (Fig. 6A-C). In control tissue, the average area of Sirius Red staining representing collagen in the CSM layer was 12.4 ± 0.7% of total area (Fig. 6D). The amount of collagen labeling was variable among achalasia biopsy samples but significantly increased overall (23.5 ± 6.3%, P < 0.001) relative to control samples (Fig. 6D), while the percent of total collagen was similar between the PE and LES of achalasia patients (not shown). However, there was no significant difference in collagen staining among tissue samples from the 3 subtypes of achalasia (Fig. 6E). We then performed a correlation analysis to determine if patient factors such as age, Eckhardt score or duration of symptoms correlated with collagen deposition. However, none of these factors appeared to correlate with an increase in collagen in the achalasia biopsies, suggesting no relationship between the examined disease characteristics and amount of collagen deposition in the CSM layer of the affected tissue.

Figure 6. Intramuscular collagen staining is significantly increased in the circular smooth muscle (CSM) layer of the esophagus in achalasia. (A-C) Representative images of histochemistry for detection of collagen using Sirius Red/Fast Green staining, in sections of control esophagus (A), type 1 achalasia (B) and type 3 achalasia (C). The type 3 pattern was typically more aberrant than the other subtypes of achalasia. Arrows indicate regions of red staining (collagen) among the smooth muscle cells (intracellular regions, green). Scale bar, 50 µm. (D) Outcome of image analyses showing increased amount of collagen in achalasia, expressed as % total area of control or biopsy CSM sections. *P < 0.01 vs control. (E) Image analysis showing that the relative amounts of collagen were similar among the individual subtypes of achalasia.

Achalasia is a disorder characterized by impaired LES relaxation and aperistalsis in the smooth muscle segment of the esophagus, resulting primarily from a loss of inhibitory myenteric neurons in the smooth muscle esophagus.⁴ To date, relatively little attention has been paid to changes in esophageal smooth muscle that may accompany the major loss of inhibitory innervation. In the current study, we obtained esophageal CSM during the minimally invasive POEM procedure, and confirmed a major deficiency of all axons innervating the CSM in achalasia. This was accompanied by an increased expression of major phenotypic smooth muscle cell markers, raising the possibility that altered smooth muscle function may contribute to the pathophysiology of achalasia.

Esophageal smooth muscle cells display a lack of innervation in patients with achalasia, which is thought to be mediated through an inflammatory process involving T lymphocytes and mast cells.^5,11,20 In inflammatory models of disease elsewhere in the gastrointestinal system, such as the 2,4,6-trinitrobenzene sulfonic acid-model of Crohn’s disease, there is damage to axonal processes as well as the loss of myenteric neurons.¹⁰ However, there is an increase in GDNF expression by the CSM, which promotes the subsequent growth of axons to re-establish innervation density and thereby restore function over time. The current data suggests that a similar phenomenon does not occur in achalasia, presumably because the inflammation-induced destruction of myenteric neurons is ongoing, rather than a transient, self-limiting injury.

In vitro studies show that neuronal NO inhibits intestinal smooth muscle growth and that damage to nitrergic innervation correlates with the onset of CSM hyperplasia and an altered phenotype in vivo.²¹ Thus, the early events of achalasia could include the compromise of NO expression, with impact on its dual roles in the regulation of smooth muscle function and phenotype. In rat models of Crohn’s-related stricturing disease, denervated smooth muscle cells tend to proliferate and lead to stricture formation.¹⁰ We hypothesized that denervated CSM in the esophagus of achalasia patients would also proliferate, thereby contributing to the increased thickness of the smooth muscle layer reported by others.²² Interestingly, there was no increase in Ki67 staining in any of the achalasia samples, regardless of duration of symptoms. This suggests either that proliferation occurs very early in the disease process or that hyperplasia is not part of the pathogenesis of achalasia. Certainly, there was a significant increase in collagen in the tissue samples and this may be an important contributor to the increased thickness of the esophageal smooth muscle.

The significant upregulation in SMA and SMOO expression in the smooth muscle of achalasia samples suggests an increased contractile phenotype, although in vitro contractility studies are needed to confirm this. The mechanism leading to this increase is unknown, but studies have shown that TGFβ signaling can contribute to upregulation of smooth muscle markers such as SMA and SM22²³ and an increase in TGF expression in achalasia has been reported.²⁴ Further, signaling through this fibrogenic cytokine has also been shown to lead to an increase in collagen²⁵ and may contribute to our observations in the esophagus of achalasia patients.

Tissue analysis using RT-PCR is a powerful way of determining the message level of a variety of key proteins. In this study, we analyzed tissue from 5 achalasia patients to determine the levels of collagen, smooth muscle contractility markers and GDNF. Despite the small sample size, the data showed an increase in SMOO, with decreases in MYH and COL3, along with possible increases in expression of SMA and COL1. While our PCR analysis had a low sample size, the outcomes were similar to a recent study²⁶ using gene expression microarray analysis on 12 LES samples removed by myotomy from achalasia patients, which found a greater than four-fold increase in SMA relative to control, as well as a greater than seven-fold increase in myosin heavy chain 11, an additional major contractile protein in smooth muscle.

GDNF is a key neurotrophin in the post-natal enteric nervous system.²⁷ GDNF is secreted in large part by gut smooth muscle cells in order to support the survival and axonal growth of enteric neurons.¹⁸ In other studies, we have observed that smooth muscle supports its own innervation by secretion of GDNF;²⁸ however in esophageal smooth muscle from achalasia patients, the apparent increase in GDNF transcript appears unable to lead to the re-establishment of innervation to the smooth muscle wall, likely because of the severe ongoing injury to the myenteric neurons.

Previous studies have compared subtypes of achalasia, noting that Types I and II are associated with fewer myenteric plexus neurons than Type III. We were unable to detect any quantitative differences in axon staining between any of the types of achalasia, although this may be a result of under-sampling for both Type I/III. Nonetheless, in our study, it appeared that the expression of markers of smooth muscle cell contractility was higher in Type I than the other types of achalasia. This may be in keeping with a belief that Type I achalasia represents a longer duration of disease compared to the other subtypes.^5,29

There are several limitations to this study. Firstly, most of the patients undergoing this procedure had Type II achalasia (the most common subtype), thereby making comparisons among the 3 groups challenging due to sampling error. Our sample size was relatively small; but despite this we were able to match data regarding the patient’s clinical presentations with their histologic findings. Similarly, we only had 3 control patients; however, they appeared similar to each other in all of our comparisons and therefore we believe the likelihood of sampling error is low. Biopsy samples were also small, which limited our ability to apply multiple tests per biopsy. As a result, the samples for PCR analysis were taken from a separate group of patients than those from which immunocytochemistry was performed. We also limited biopsies to the CSM in order to minimize the risk of perforation through the deeper longitudinal muscle and serosa. As a result we were unable to visualize the myenteric plexus as others have reported using this technique.¹¹ However, myenteric plexus pathology has been well documented previously using full thickness samples obtained surgically.

In conclusion, we present, for the first time, direct evidence that the axonal innervation of esophageal smooth muscle is significantly impaired in patients with achalasia. Further, achalasia is characterized by altered smooth muscle cell phenotype and increased intramuscular collagen deposition. Importantly, these novel findings were discovered by using specimens taken directly from the CSM in patients undergoing POEM and supports an ongoing role for this technique in further exploring the pathophysiology of achalasia.

Note: To access the supplementary tables mentioned in this article, visit the online version of Journal of Neurogastroenterology and Motility at http://www.jnmjournal.org/, and at https://doi.org/10.5056/jnm23024.

The authors express their appreciation to Drs Wiley Chung and David Hurlbut for assistance in obtaining control tissue specimens.

This study was supported by a Research Award through the Department of Medicine, Queen’s University (to Robert Bechara).

None.

Robert Bechara, William G Paterson, Michael G Blennerhassett, and David M Rodrigues conceived and planned the study; Robert Bechara collected all tissue samples; David M Rodrigues, Michael G Blennerhassett, Sandra R Lourenssen, and Jay Kataria conducted analysis and data interpretation; and David M Rodrigues, Michael G Blennerhassett, and Sandra R Lourenssen drafted the manuscript. All authors assisted in editing.