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. 2024 Oct 3;72(10):611–622. doi: 10.1369/00221554241287267

Intestinal Tuft Cells Are Enriched With Protocadherins

Rachel Stubler 1, Sarah A Dooley 2, Rachel Edens 3, Maribeth R Nicholson 4, Amy C Engevik 5,✉
PMCID: PMC11471013  PMID: 39360911

Abstract

Intestinal tuft cells are rare cells that regulate diverse functions. They harbor chemosensory receptors and signal to the mucosal immune system in response to external stimuli, though their full function and structure remain unclear. Named for their apical “tuft” of long actin-rich microvilli, tuft cells facilitate chemoreception and other physiological responses. In enterocytes, microvilli are stabilized by intermicrovillar adhesion complexes (IMACs) composed of several proteins, including cadherin-related family member-2 (CDHR2) and cadherin-related family member-5 (CDHR5), Myosin 7b, and Usher syndrome type 1 C (USH1C). We hypothesized that IMACs would be enriched in tuft cells to regulate microvillar organization. Immunostaining of murine intestinal tissue revealed that CDHR2 and CDHR5 colocalize with the tuft cell markers, DCLK1, phospho-EGFR, advillin, and cytokeratin 18. CDHR2 was dispersed throughout murine tuft cells, while CDHR5 was concentrated on the apical surface. USH1C and Myosin 7b were present in tuft cells, but at lower levels. Human single-cell RNA sequencing revealed robust CDHR2 and CDHR5 expression in tuft cells in the small intestine and colon. Immunostaining of human intestinal tissue confirmed CDHR2 and CDHR5 localization to the apical surface of tuft cells. Our findings demonstrate that protocadherins are key components of murine and human intestinal tuft cells.

Keywords: epithelium, gut, IMAC, microvilli

Introduction

The intestinal epithelium forms a protective barrier that facilitates nutrient absorption and excludes harmful luminal contents and bacteria from entering the body. The epithelium is composed of diverse cell types including absorptive enterocytes, mucin-secreting goblet cells, proliferative stem cells, hormone-producing enteroendocrine cells, and rare chemosensory cells known as tuft cells. While tuft cells were identified in the small intestine and colon in the 1970s,1–3 their structure and function in the intestinal epithelium remains incompletely understood. Tuft cells harbor type 1 and type 2 taste receptors and succinate receptors allowing them to sample the luminal environment and elicit signaling cascades to trigger immune- and neuronal-mediated responses.4–6

Interest in tuft cells has increased since the publication of multiple studies showing that intestinal tuft cell expansion is induced by protist and helminth infection through activation of group 2 innate lymphoid cells (ILC2s).6–8 These studies have been facilitated in part by the development of improved antibodies for tuft cell identification. Many proteins have been identified as tuft cell markers. One of the most common markers of tuft cells is doublecortin-like kinase 1 (DCLK1), a kinase that functions in microtubule polymerization.9–11 Other structural markers include the intermediate filament cytokeratin 18 (CK-18), acetylated α-tubulin (Ac-TUBA), and the actin-bundling protein, advillin (AVIL).10,12–14 In addition, tuft cells specifically express the transcription factor, POU class 2 homeobox 3 (POU2F3), 8 and are enriched for EGFR phosphotyrosine 1068 (P-EGFR). 15 While not specific for tuft cells, tuft cells can also be identified by immunostaining based on their enrichment of actin at the apical membrane. 16

Tuft cells are named for their apical “tuft” of long actin-rich protrusions known as microvilli. There are many differences between microvilli in tuft cells and those in neighboring absorptive enterocytes. In enterocytes, microvilli are remarkably uniform in length and width and are tightly packed into a hexagonal arrangement. 17 Each microvilli contains approximately 30–40 actin filaments which are anchored to the terminal web, a dense actomyosin and intermediate filament-rich subapical structure. 18 In contrast, tuft cell microvilli are much thicker, consisting of around 100 actin filaments intertwined with acetylated microtubules. 19 The actin and microtubule network protrudes deep into the tuft cell body, which lacks an anchoring terminal web. 20 Although these microvilli are much wider and longer, they maintain hexagonal packing. 19

In enterocytes, hexagonal packing is regulated by the intermicrovillar adhesion complex (IMAC), a protein complex which links neighboring microvilli at their distal tips. Cadherin-related family member-2 [CDHR2, also known as protocadherin-24 (PCDH24)] and cadherin-related family member-5 [CDHR5, also known as mucin-like protocadherin (MLPCDH)] form adhesion linkages in the intermicrovillar space. 17 The cytoplasmic domains of these protocadherins bind to the scaffolding proteins ankyrin repeat protein and sterile a motif domain containing 4B (ANKS4B) and Usher syndrome 1C (USH1C).21,22 These IMAC components are trafficked to the microvillar tips and anchored to the actin cytoskeleton by the motor protein, myosin 7b (MYO7B). 23 Loss of IMAC components in enterocytes results in microvillar dysregulation and is thought to impact susceptibility to inflammation. 24

While it is known that tuft cell microvilli differ from enterocyte microvilli, 16 few studies have investigated how these tuft cell microvilli are organized and whether IMAC proteins are uniquely enriched in tuft cells. To determine whether murine and human tuft cells express CDHR2, CDHR5, and the other IMAC proteins, we performed immunostaining for IMAC proteins alongside an assortment of tuft cell markers. We found that CDHR2 and CDHR5 are enriched in tuft cells compared with neighboring cells, while MYO7B and USH1C are expressed in similar amounts in tuft cells and absorptive cells in the small intestine and colon. CDHR2 and CDHR5 localize to the apical surface with some subapical staining in mouse tissue, while only apical CDHR2 and CDHR5 are present in human tuft cells. Collectively, these data identify that protocadherins are highly expressed in intestinal tuft cells.

Materials and Methods

Animal and Tissue Collection

The Institutional Animal Care and Use Committee of the Medical University of South Carolina approved all protocols for animal care and maintenance. C57BL/6J mice were obtained from the Jackson Laboratory. Adult male and female mice were used in all experiments to account for any sex differences that may affect the expression of cell lineage markers and IMAC components. Intestinal tissue was harvested from adult mice and fixed in 10% neutral-buffered formalin at 4C overnight. The tissue was then processed and paraffin-embedded.

Human Samples

Ileal and colonic biopsy specimens were obtained from archived pediatric patient specimens (≤18 years old) at Monroe Carell Junior Children’s Hospital at Vanderbilt (MCJCHV). Healthy control patients were defined as patients without a diagnosis of inflammatory bowel disease who underwent endoscopy and were found to have normal pathology. All subjects previously consented to human tissue acquisition, storage, and research, and the study was approved by the institutional review board. Specimens were fixed in neutral-buffered formalin and paraffin-embedded as part of the standard pathology procedures.

Staining

Slides containing 5 µm paraffin tissue sections were baked at 60C for 15 min to promote tissue adherence. Samples were deparaffinized using Histoclear (HS200; National Diagnostics, Atlanta, GA) and rehydrated through a series of graded ethanol washes. In a pressure cooker, antigen retrieval was performed in citrate buffer (pH 6) or tris buffer (pH 9) to expose epitopes. Slides were cooled on ice, washed in phosphate-buffered saline (PBS), and blocked with Dako serum-free protein block (X0909; Dako, Santa Clara, CA) for 90 min at room temperature in a humidified chamber. Mouse on mouse block (MKB-2213; Vector Laboratories, Newark, CA) was applied for 15 min for samples being stained with unconjugated mouse antibodies. Primary antibodies were diluted in Dako antibody diluent with background-reducing components (S3022; Dako) and incubated overnight at 4C in a humidified chamber. The following day, slides were washed in PBS three times followed by the addition of secondary antibodies diluted 1:200 in Dako antibody diluent (S0809; Dako). Tissue sections were incubated for 1 hr at room temperature in a humidified chamber. Hoescht (62249, 12.3 mg/ml; Thermo Fisher Scientific, Waltham, MA) was diluted 1:1000 in PBS and added for 5 min. Finally, slides were coverslipped with Prolong Gold Antifade (P36934; Thermo Fisher Scientific) and allowed to dry before imaging. Immunohistochemistry was performed using IHC Prep and Detect Kits according to manufacturer’s protocol (PK10017 and PK10018; Proteintech, Rosemont, IL).

Primary and Secondary Antibodies

Primary antibodies and dilutions used in this study are as follows: mouse γ-actin 1:100 (sc-65638; Santa Cruz Biotechnology, Dallas, TX); rabbit CDHR5 1:500 (HPA009081; Sigma, Burlington, MA); mouse microprotocadherin/CDHR5 1:100 (sc-16953; Santa Cruz Biotechnology); rabbit PCDH24/CDHR2 1:200 (27103-1-AP; Proteintech); rabbit CDHR2 1:400 (HPA1010569; Sigma); mouse DCLK1 1:200 (MA5-26800; Invitrogen, Waltham, MA); rabbit DCLK1 1:200 (ab109029; abcam, Waltham, MA); rabbit advillin 1:200 (NBP2-34118; Novus, Centennial, CO); rabbit phospho-EGFR 1:100 (ab205828; abcam); mouse cytokeratin18 1:100 (NBP2-47984AF594; Novus); and rabbit POU2F3 1:200 (HPA019652; Sigma). Donkey secondary antibodies conjugated to Alexa Fluor 488, Cy3, Alexa Fluor 594, Alexa Fluor 647, or Cy5 were diluted 1:200 (Jackson ImmunoResearch, West Grove, PA).

Imaging

Micrographs were obtained from a Leica SP8 Microscope (Wetzlar, Germany) using the 63× oil immersion objective and a Zeiss Axio Imager.M2 Microscope (Oberkochen, Germany) using the 20× objective. Single-channel grayscale tiff images were exported using Leica and Zen software and merged into red, green, and blue using Adobe Photoshop. Immunohistochemistry slides were imaged using the 20× objective on an Olympus BX40 microscope (Tokyo, Japan).

Intestinal Organoids

Crypts isolated from the small intestine of C57BL/6J mice were used for organoid generation. Organoids were grown in expansion media for 6–9 days and then differentiated for 48–72 hr. To drive the development of tuft cells, differentiation media was supplemented with 400 ng/ml recombinant murine IL-4 (214-14; Peprotech, Cranbury, NJ) and 400 ng/ml recombinant murine IL-13 (210-13; Peprotech).8,25 Organoids were fixed with 4% paraformaldehyde for 30 min at room temperature and then processed for paraffin-embedding.

Results

Intestinal Tuft Cells Display an Enrichment of Protocadherins

To assess the distribution of CDHR2, we performed immunohistochemical staining of C57BL6/J mouse intestine. As expected, CDHR2 was highly expressed on the apical membrane of absorptive enterocytes. Interestingly, cells with a bottle-shaped tuft cell morphology displayed diffuse cytoplasmic staining for CDHR2 (Fig. 1A). To determine whether these cells were in fact tuft cells, we immunostained the mouse intestine with the tuft cell markers DCLK1, CK-18, and Ac-TUBA as well as CDHR2. We found that CDHR2 colocalized with the tuft cell markers DCLK1, CK-18, and Ac-TUBA in both the small intestine and colon (Fig. 1B to D). Interestingly, CDHR2 localized throughout the cell body but was enriched at the apical surface of the cell, colocalizing strongly with the apical membrane marker γ-actin. Quantification of 20 images from four mice revealed that CDHR2 was expressed by 93% of DCLK1-expressing tuft cells (Fig. 3A). The majority of DCLK1-expressing tuft cells that did not co-stain for CDHR2 were localized to the crypt base, suggesting that CDHR2 expression may be more important in tuft cells which are in closer contact with the luminal contents.

Figure 1.

Figure 1.

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CDHR2 is highly expressed in murine tuft cells. (A) Immunohistochemical staining of CDHR2 in the small intestine, highlighting cells that express CDHR2 throughout the cell body. (B-D) Immunofluorescent staining of small intestine and colon for CDHR2 (magenta), apical membrane marker γ-actin (yellow), tuft cell markers (green): DCLK1 (B), CK-18 (C), and Ac-TUBA (D). Scale bar = 50 µm (A low magnification), 5 µm (A Inset, B-D).

Figure 3.

Figure 3.

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CDHR2 and CDHR5 are expressed by a high percentage of murine tuft cells. (A) Immunostaining of CDHR2 (magenta), DCLK1 (green), and γ-actin (yellow) with quantification of the percentage of DCLK1-positive cells that co-express CDHR2. (B) Immunostaining of CDHR5 (magenta), DCLK1 (green), and γ-actin (yellow) with quantification of the percentage of DCLK1-positive cells that co-express CDHR5 and γ-actin. n=4 mice/group. Scale bar = 50 µm.

CDHR5 is another protocadherin highly expressed by the intestinal epithelium. In line with other studies, we found by immunohistochemistry staining that CDHR5 was present at high levels on the apical membrane. Similar to our CDHR2 staining, we found that some cells that resembled tuft cells displayed an enrichment of CDHR5 at or just below the apical membrane (Fig. 2A). Immunofluorescent staining with a panel of tuft cell markers confirmed that the cells that expressed high levels of CDHR5 were tuft cells. We found that CDHR5 was highly expressed at the apical membrane with some subapical staining in murine tuft cells, marked by DCLK1, POU2F3, γ-actin, AVIL, and CK-18 (Fig. 2B to D). Quantification revealed that CDHR5 was present in 45% of DCLK1-expressing tuft cells (Fig. 3B). Because CDHR5 is largely localized to the apical membrane, we quantified CDHR5 in DCLK1-positive tuft cells in which the tuft was visible by γ-actin staining. CDHR5 staining was present in 87% of γ-actin and DCLK1-positive tuft cells. In addition, 6% of tuft cells did not stain for γ-actin but showed subapical CDHR5 staining. Taken together, these data indicate that CDHR5 expression is highly conserved in murine tuft cells. To compare the localization of different protocadherins in tuft cells, immunostaining was performed with CDHR2, CDHR5, and tuft cell marker P-EGFR (Fig. 4). CDHR2 and CDHR5 colocalized at the apical surface of tuft cells in the small intestine and colon. CDHR2 displayed more cytoplasmic staining than CDHR5, and subapical CDHR5 did not colocalize with CDHR2. These data suggest that tuft cells are highly enriched for protocadherins CDHR2 and CDHR5.

Figure 2.

Figure 2.

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CDHR5 localizes to the apical surface of murine tuft cells. (A) Immunohistochemical staining of CDHR5 in the small intestine, highlighting enrichment of CDHR5 in a select cell. (B-D) Immunofluorescent staining of murine small intestine and colon for CDHR5 (magenta) with tuft markers: (B) DCLK1 (green) and apical marker γ-actin (yellow), (C) POU2F3 (green) and γ-actin (yellow), and (D) AVIL (green) and CK-18 (yellow). Scale bar = 50 µm (A low magnification), 5 µm (A Inset, B-D).

Figure 4.

Figure 4.

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CDHR2 and CDHR5 display differential staining patterns in tuft cells. Immunostaining of murine small intestine and colon showing that CDHR2 (magenta) and CDHR5 (green) are both enriched at the apical surface of tuft cells, marked by P-EGFR (yellow). CDHR2 displays diffuse more cytoplasmic staining compared with CDHR5. Scale bar = 5 µm.

USH1C and Myosin 7b Are Moderately Expressed by Tuft Cells

As protocadherins are known to be major components of the IMACs which regulate microvillar packing in enterocytes, we sought to determine whether other IMAC components were expressed by tuft cells. To address this question, we performed immunofluorescent staining of Myosin 7b, the molecular motor that traffics IMAC components and anchors the IMAC to the actin cytoskeleton, and USH1C, the protein that links Myosin 7b and the protocadherins, CDHR2 and CDHR5. While Myosin 7b and USH1C were present in tuft cells in the small intestine and colon, they did not appear significantly enriched compared with neighboring cells (Fig. 5). This suggests that tuft cells harbor the components of IMACs, with a unique enrichment of protocadherins.

Figure 5.

Figure 5.

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Other IMAC components are not enriched in tuft cells compared with neighboring enterocytes. Immunostaining of murine small intestine and colon for (A) MYO7B (magenta) and (B) USH1C (magenta) with DCLK1 (green) and γ-actin (yellow). Scale bar = 5 µm.

Tuft Cells in Intestinal Organoids Maintain High Expression of Protocadherins

Many factors are known to impact tuft cells, including the gut microbiota, immune cells, and the enteric nervous system.26–29 To confirm that tuft cells express protocadherins independently of these external signals, we generated intestinal organoids from C57BL6/J mice. Organoids were grown in 3D in Matrigel and differentiated for 48 hr in differentiation media supplemented with 400 ng/ml IL-4 and 400 ng/ml IL-13 to promote tuft cell differentiation. Immunostaining of protocadherins with various tuft cell markers confirmed that CDHR2 strongly stains the apical surface of tuft cells with diffuse staining in the cell body, while CDHR5 localizes to the apical surface with some subapical staining (Fig. 6). These data suggest that tuft cells, independent of other signaling factors, endogenously express high levels of protocadherins.

Figure 6.

Figure 6.

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Tuft cells in murine intestinal organoids are enriched for CDHR2 and CDHR5 and display staining patterns consistent with murine tissue. Immunostaining for: (A) CDHR2 (magenta) with DCLK1 (green) and γ-actin (yellow), (B) CDHR5 (magenta) with DCLK1 (green) and γ-actin (yellow), (C) CDHR2 (magenta) with CK-18 (green) and γ-actin (yellow), (D) CDHR5 (magenta) with AVIL (green) and CK-18 (yellow), (E) CDHR2 (magenta) with P-EGFR (green) and Ac-TUBA (yellow), (F) CDHR5 (magenta) with POU2F3 (green) and γ-actin (yellow), and (G) CDHR2 (magenta) and CDHR5 (green) with P-EGFR (yellow). Scale bar = 10 µm on low magnification and 5 µm on high magnification insets.

Human Tuft Cells in the Small Intestine and Colon Express Protocadherins at the Apical Membrane

To determine whether human tuft cells in the small intestine and colon express IMAC components, we queried CellXGene, a publicly available single-cell RNA sequencing database housing more than 1000 datasets. Consistent with our findings in murine tissue, human intestinal tuft cells exhibited high levels of CDHR2 and CDHR5 co-expressed with the known tuft cell marker POU2F3 (Fig. 7A). Interestingly, CDHR5 was in the top 60 genes expressed by human tuft cells according to publicly available single-cell sequencing data through CellXGene. 30 We next sought to confirm the levels of IMACs by immunostaining. Staining of human small intestine and colon identified that CDHR2 and CDHR5 are also present in human tuft cells (Fig. 7B and C). We observed that CDHR2 and CDHR5 colocalized with γ-actin at the apical surface of tuft cells in human tissue. Unlike murine tuft cells, which displayed subapical staining of both CDHR2 and CDHR5, human tuft cells displayed little to no subapical staining. Together, these findings suggest that protocadherins are a previously unrecognized component of intestinal tuft cells in mice and humans and may play a role in tuft cell function.

Figure 7.

Figure 7.

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CDHR2 and CDHR5 localize to the apical surface of tuft cells in the human small intestine and colon. (A) Single cell RNA sequencing of tuft cells in healthy control human small intestine and colon from CellXGene publicly available database. (B-C) Immunostaining of human small intestine and colon for (B) CDHR2 (magenta) and (C) CDHR5 (magenta) with P-EGFR (green) and γ-actin (yellow). Scale bar = 10 µm on low magnification and 2 µm on high magnification insets.

Discussion

In this study, we identified that intestinal tuft cells display enrichment for the protocadherins, CDHR2 and CDHR5. In murine tissue and mouse intestinal organoids, CDHR2 and CDHR5 were enriched at the apical membrane and displayed differential patterns of subapical staining. In contrast, human intestinal tuft cells contained CDHR2 and CDHR5 exclusively localized to the apical membrane. To the best of our knowledge, this is the first observation of protocadherin enrichment in murine and human tuft cells. Our data suggest that CDHR2 and CDHR5 may play unique roles in intestinal tuft cells. We postulate that the primary function of apical CDHR2 and CDHR5 is adhesion to regulate the organization and stabilization of tuft cell microvilli.

One of the interesting observations from our work was that although both murine and human tuft cells had enrichment of protocadherins; they differed in terms of their localization within the cells. The differences in staining patterns between mouse and human tuft cells could be due to species-dependent differences in the protein structure. CDHR2 and CDHR5 are canonically thought to form heterophilic linkages between neighboring microvilli in enterocytes. Gray et al. 31 recently published a study identifying that mouse and human CDHR2 and CDHR5 have poor sequence and structural conservation. The authors found that CDHR2 is able to form homophilic adhesive linkages in humans, while CDHR5 is able to form homophilic adhesive linkages in mice. 31 CDHR2 may be binding to itself in murine tuft cells creating the diffuse staining pattern. In murine tissue, CDHR5 mainly localized to the apical membrane, with some staining seen directly below the apical membrane. This subapical staining pattern was recently reported in a preprint by Silverman et al. 19 who postulated that CDHR5 may be localized in vesicles being transported along microtubules to the apical surface. The staining pattern visualized would be consistent with cellular trafficking to the apical surface. The roles of cytoplasmic CDHR2 and CDHR5 should be investigated in subsequent studies.

The protocadherin family is a family of adhesion proteins that are highly expressed in neural tissues. 32 In neural tissues, the extracellular domains of protocadherins are known to play key roles in cell–cell adhesion, while the intracellular domains, which display low to moderate sequence homology, participate in a variety of intracellular signaling cascades. 33 The intestinal protocadherins, CDHR2 and CDHR5, have been well studied for their adhesion functions, but far fewer studies have investigated the roles of their intracellular domains in cell signaling. A few studies have suggested roles of CDHR2 and CDHR5 in regulating WNT signaling in colon cancer cell lines.34–36 CDHR2 downregulates WNT signaling via galectin-1 and galectin-3 interactions with its intracellular domain. These interactions sequester β-catenin and lead to PI3K inactivation. 36 CDHR5 also plays a role in limiting β-catenin transcriptional activity in cell lines. Similar to CDHR2, the cytoplasmic domain of CDHR5 is able to sequester β-catenin.37,38 In addition, the extracellular domain is able to reduce WNT signaling through opposition of EGFR and AKT signaling which decreases β-catenin stability. 37 It is possible that the intracellular domain of CDHR2 and the intracellular and extracellular domains of CDHR5 could be involved in WNT signaling cascades in intestinal tuft cells, thereby controlling intestinal response to a variety of external stimuli.

Our data suggest that the packing of tuft cell microvilli may be regulated in part by CDHR2 and CDHR5, which were enriched at the apical surface of mouse and human tuft cells. However, we did not observe tuft cell enrichment for other enterocyte IMAC components such as MYO7B and USH1C. We postulate that CDHR2 and CDHR5 may require different scaffolding proteins in tuft cells due to the differences in microvillar structure. It is possible that CDHR2 and CDHR5 could be anchored to microtubules rather than actin filaments in these cells.

A limitation of this study is the sole use of immunostaining as an approach to investigate tuft cells. Due to the rarity of these cells, it is not feasible to quantify protein levels in tuft cells using current cell sorting-based techniques. In addition, this study did not investigate the mechanisms underlying CDHR2 and CDHR5 localization in intestinal tuft cells. Future studies are needed to elucidate the role of subapical CDHR2 and CDHR5 and their binding partners at the apical membrane in intestinal tuft cells.

In summary, this study identifies CDHR2 and CDHR5 as previously unrecognized components of intestinal tuft cells in mice and humans. CDHR2 and CDHR5 were enriched at the apical membrane, suggesting that they may play a role in organizing tuft cell microvillar packing and may regulate tuft cell physiology.

Acknowledgments

We would like to acknowledge the MUSC DDRCC imaging core for the use of their microscopes.

Footnotes

The author(s) declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Author Contributions: Conceptualization: RS and ACE; methodology and data acquisition: RS and ACE; manuscript preparation RS; review and editing: RS, SAD, RE, MRN, and ACE; funding acquisition: RS, SAD, RE, and ACE. All authors have read and agreed to the published version of the manuscript.

Funding: The author(s) disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: This study was supported by the National Institutes of Health (NIH) grant K01 DK121869 to ACE, startup funds from MUSC to ACE, and MUSC’s P30 DK123704 and P20 GM120475. This work was also supported in part by TL1 TR001451, UL1 TR001450, and the HCS cornerstone grant to RS and DK139736 to SAD and GM132055 to RE.

Contributor Information

Rachel Stubler, Department of Regenerative Medicine & Cell Biology, Medical University of South Carolina, Charleston, SC.

Sarah A. Dooley, Department of Regenerative Medicine & Cell Biology, Medical University of South Carolina, Charleston, SC

Rachel Edens, Department of Regenerative Medicine & Cell Biology, Medical University of South Carolina, Charleston, SC.

Maribeth R. Nicholson, Division of Pediatric Gastroenterology, Hepatology and Nutrition, Monroe Carell Jr. Children’s Hospital at Vanderbilt, Nashville, TN

Amy C. Engevik, Department of Regenerative Medicine & Cell Biology, Medical University of South Carolina, Charleston, SC.

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