Elucidating the Mechanism Behind Gastric Restitution

A dissertation submitted to the Graduate School of the University of Cincinnati in partial fulfillment of the requirements for the degree of

Doctor of Philosophy

in the Department of Pharmacology and Systems Physiology of the College of Medicine by

Kristen Anette Engevik B.S. 2012 Biola University, La Mirada, California

Committee Chair: Marshall H. Montrose, Ph.D.

Committee Members: Judith Heiny, Ph.D.; Anjaparavanda Naren, Ph.D.; Yana Zavros, Ph.D.; Tongli Zhang, Ph.D. Abstract

Background: The gastric mucosa of the stomach is continually exposed to environmental and physiological stress factors which can cause local epithelial damage.

While much is known about the complex nature of gastric wound repair, the stepwise process that characterizes epithelial restitution remains poorly defined.

Objective: This work seeks to elucidate effectors that drive gastric epithelial repair using a reductionist culture model, gastric organoids.

Major Findings:

Chapter 2 Assessing permeability, repair and cell death in the gastric organoid system

This work establishes approaches to assess restitution in gastric organoids as well as identify the type of cell death induced by two photon microscopy. Two photon-induced photodamage results in caspase-activated apoptosis in the damaged cell, which mirrors natural cell shedding within gastric organoids. Similar to in vivo, localized photodamage results in rapid dead cell exfoliation, coinciding with the migration of adjacent viable cells to cover the denuded area, sustaining epithelial continuity. Measurement of dead cell exfoliation and closure of the damage area are reliable analyses to assess epithelial repair in gastric organoids. Under normal conditions, Lucifer yellow serves as a consistent extracellular marker to indicate epithelial barrier integrity. However in deficiency conditions, Lucifer yellow does not yield quantitative data to shed more light upon gastric organoid repair.

Chapter 3 Comparison of genetically encoded calcium sensors to assess calcium mobilization in gastric organoid repair studies

While calcium is a known accelerator for gastric repair, prior studies have been limited in the available technology for measuring calcium signaling. With the recent generation of a more sensitive genetically encoded calcium indicator (GECI) ( transgenic mouse, we tested organoids from mice expressing the previously generated GECI Yellow

Cameleon 3.0 (YC 3.0) and a modified Yellow Cameleon Nano-15 (YC Nano) which has been reported to have improved sensitivityto determine the most consistent and sensitive calcium reporter. While both YC Nano and YC 3.0 organoids exhibited similar organoid size and response to damage, imaging outcomes suggest YC Nano as a more sensitive and reliable calcium sensor.

Chapter 4 Trefoil Factor 2 activation of CXCR4 requires calcium mobilization to drive epithelial repair in gastric organoids

This work also elucidates the signaling cascade of epithelial repair using murine gastric organoids, allowing us to define which regulatory processes are intrinsic to epithelial cells. Additionally, this research allows for the validation and dissection of the signaling cascade with more precision than is possible in vivo. Following single cell damage, intracellular calcium selectively increases within cells adjacent to the damage site and is essential to promote repair. Our findings demonstrate TFF2 acts via CXCR4 and EGFR signaling, including ERK activation, to drive Ca2+ mobilization and promote gastric repair. Sodium hydrogen exchanger 2, while essential for repair, acts downstream of

TFF2 and calcium mobilization.

Chapter 5 Sources of localized calcium mobilization during gastric organoid epithelial repair

This work seeks to identify potential sources of calcium mobilization during repair.

Based upon published RNA sequence data, several highly expressed calcium associated genes were chosen to target calcium mobilization. Inhibition studies suggest that extracellular sources via voltage gated calcium channels are essential for calcium mobilization and proper repair. Furthermore, calcium mobilization via phospholipase C signaling releases calcium stores from the ER, which are required to promote repair.

These studies suggest both extracellular and intracellular sources are essential for calcium dependent epithelial restitution.

Acknowledgements

I would like to thank my advisor, Dr. Marshall Montrose for his guidance and thoroughness during my graduate work. I would also like to thank the members of the

Montrose lab for their assistance with experimental design and execution as well as for their scientific rigor. A special thanks to my sisters, Mindy and Amy, for the unwavering support, encouragement, and input they’ve given me throughout my Ph.D. candidacy. Table of Contents Abbreviations ...... 1 Chapter 1 Literature Review ...... 3 1.1 Gastric physiology and anatomy ...... 4 1.2 Disruption of barrier function ...... 7 1.3 Gastric epithelial restitution ...... 9 1.4 Trefoil factor family peptides ...... 11 1.5 Epithelial Growth Factors in gastric restitution ...... 14 1.6 Sodium hydrogen exchangers in gastric restitution ...... 16 1.7 Role of calcium in repair ...... 17 1.8 Genetically encoded calcium indicators ...... 1 1.9 Gastric cancer lines vs. gastric organoids ...... 21 1.10 Major Contributions ...... 24 Chapter 2 Assessing permeability, repair and cell death in the gastric organoid system ...... 25 2.1 Abstract ...... 26 2.2 Introduction ...... 27 2.3 Materials and Methods ...... 29 2.3.1 Animal husbandry ...... 29 2.3.2 Mouse-derived corpus organoid culture...... 29 2.3.3 Induction of two-photon laser-induced photodamage ...... 30 2.3.4 Microinjection ...... 31 2.3.5 Image analysis ...... 31 2.3.6 Statistical analysis ...... 32 2.4 Results ...... 33 2.4.1 Assessing organoid permeability using Lucifer Yellow ...... 33 2.4.2 Assessing repair in various organoid models ...... 36 2.4.3 Assessing repair in TFF2 deficient gastric organoids ...... 38 2.4.4 Photodamage induces caspase-activated apoptosis ...... 39 2.5 Discussion ...... 43 Chapter 3 Comparison of genetically encoded calcium sensors to assess calcium mobilization in gastric organoid repair studies ...... 47 3.1 Abstract ...... 48 3.2 Introduction ...... 49 3.3 Materials and Methods ...... 53 3.3.1 Animal husbandry ...... 53 3.3.2 Mouse-derived corpus organoid culture...... 53 3.3.3 Induction of two-photon laser-induced photodamage ...... 54 3.3.4 Image analysis ...... 54 3.3.5 Statistical analysis ...... 55 3.4 Results ...... 56 3.4.1 Organoid size of YC 3.0 and YC Nano is consistent with other organoid models ...... 56 3.4.2 Quality of imaging and date from YC 3.0 and YC Nano ...... 56 3.4.3 Comparison of response to damage in YC 3.0 and YC Nano ...... 59 3.4.4 Calcium mobilization response of YC 3.0 and YC Nano ...... 60 3.5 Discussion ...... 62 Chapter 4 Trefoil factor 2 activation of CXCR4 requires calcium mobilization to drive epithelial repair in gastric organoids ...... 66 4.1 Abstract ...... 67 4.2 Introduction ...... 68 4.3 Materials and Methods ...... 71 4.3.1 Animal husbandry ...... 71 4.3.2 Mouse-derived corpus organoid culture...... 72 4.3.3 Induction of two-photon laser-induced photodamage ...... 72 4.3.4 Microinjection ...... 74 4.3.5 Image analysis ...... 75 4.3.6 Statistical analysis ...... 76 4.4 Results ...... 76 4.4.1 Organoids as a model of gastric restitution ...... 76 4.4.2 Calcium is required for epithelial wound repair in gastric organoids ...... 77 4.4.3 TFF2 receptor CXCR4 acts upstream of calcium mobilization and is involved in gastric restitution ...... 80 4.4.4 TFF2 action requires CXCR4 and calcium mobilization acting downstream during gastric restitution ...... 82 4.4.5 EGFR acts upstream of calcium mobilization and is involved in gastric restitution ...... 83 4.4.6 ERK1/2 is necessary for the repair process, acting upstream of calcium mobilization ...... 86 4.4.7 Sodium hydrogen exchanger 2 acts downstream of calcium mobilization in TFF2 driven repair ...... 87 4.4.8 Confirmation of efficacy of inhibitor concentrations used ...... 89 4.5 Discussion ...... 91 Chapter 5 Sources of localized calcium mobilization during gastric organoid epithelial repair ...... 98 5.1 Abstract ...... 99 5.2 Introduction ...... 100 5.3 Materials and Methods ...... 102 5.3.1 Animal husbandry ...... 102 5.3.2 Mouse-derived corpus organoid culture...... 103 5.3.3 Induction of two-photon laser-induced photodamage ...... 103 5.3.4 Image analysis ...... 104 5.3.5 Bioinformatics RNA sequencing data analysis ...... 105

5.3.6 Statistical analysis ...... 106 5.4 Results ...... 106 5.4.1 Identifying potential calcium targets in gastric organoids ...... 106 5.4.2 Voltage gated calcium channels are essential for intracellular calcium mobilization during repair ...... 109 5.4.3 Phospholipase C pathway is necessary for intracellular calcium mobilization during repair ...... 110 5.4.4 Store operated calcium entry is essential for calcium mobilization during restitution ...... 112 5.5 Discussion ...... 113 Chapter 6 Discussion ...... 116 6.1 Significance of Outcomes ...... 117 6.2 Future Directions ...... 123 Chapter 7 Bibliography ...... 127

Abbreviations

AGS Human gastric cancer cells ANOVA Analysis of Variation Ca2+ Calcium CXCR4 C-X-C chemokine receptor 4 CFP Cyan Fluorescent Protein DAG Diacylglycerol ddH2O Double distilled water DMSO Dimethyl sulfoxide dPBS Dulbecco's phosphate buffered saline EGF Epidermal Growth Factor EGFR Epidermal Growth Factor receptor FRET Förster resonance energy transfer GFP Green Fluorescent Protein GI Gastrointestinal GCPR G-coupled protein receptor GSII Griffonia simplicifolia lectin II g Grams H. pylori Helicobacter pylori Hoe 694 Hoechst 694

IP3 Inositol trisphosphate

IP3R Inositol trisphosphate receptor KO Knockout L Liters MLC Myosin light chain mL Milliliters mM Millimolar M Molar NHE2 Sodium Hydrogen Exchanger 2 nL Nanoliters

nm Nanometers nM Nanomolar PBS Phosphate buffered saline PD Photodamage

PIP2 Phosphatidylinositol 4,5-bisphosphate PKC Protein Kinase C PLC Phospholipase C rTFF2 Recombinant Trefoil Factor 2 ROI Region of interest RYR Ryanodine receptor SPEM Spasmolytic polypeptide expressing metaplasia SOCE Store operated calcium entry TFF1 Trefoil Factor Family 1/pS2 TFF2 Trefoil Factor Family 2/spasmolytic polypeptide TFF3 Trefoil Factor Family 3/intestinal trefoil factor Ti-Sa Titanium sapphire WT Wildtype YC Yellow Cameleon YFP Yellow Fluorescent Protein µL Microliter µm Micrometer/microns µM Micromolar

Chapter 1

Literature Review

Chapter 1 Literature Review

1.1 Gastric physiology and anatomy

The stomach is a component of the gastrointestinal (GI) tract, residing between the

esophagus and duodenum of the small intestine. As the primary site of physical

digestion of food, the stomach is a

unique organ that creates an acidic

environment to physically digest

luminal contents. The human stomach

is divided into four regions: cardia,

fundus, corpus, and antrum (Figure

1.1). The cardia is the portion that Fundus Fore- surrounds the cardioesophageal Cardia Stomach junction, or the opening of the Corpus Corpus/ Fundus esophagus into the stomach. The

Antrum fundus is the upper portion of the

stomach, while the corpus, also Figure 1.1 Anatomy of human and mouse stomach. In humans, the stomach contains anatomically distinct regions: cardia, fundus, corpus, and antrum. In mice, the stomach is divided into forestomach, known as the body, is the largest and corpus/fundus, and antrum. central part of the stomach. Finally,

the antrum is the lower part of the stomach. In mice, the stomach is divided into three

regions: the forestomach, corpus and antrum (Figure 1.1). From the lumen, beneath the

stomach mucosa lie the submucosa, outer mucosa and the inner submucosa,

muscularis externa and serosa (Figure 1.2).

Stomach

Submucosa Mucosa Muscle Layer

Serosa

Figure 1.2 Anatomy of the stomach corpus/fundus. The corpus/fundus of the stomach (from the lumen to muscle layer) consists of the mucosa, submucosa, muscle layer and serosa. In the stomach, the epithelium of the mucosa layer acts as the primary barrier against the noxious contents of the lumen. The acidity of the stomach is a necessary defense mechanism to combat harmful factors, especially pathogenic microbes. Parietal cells within the gastric epithelium are responsible for the secretion of acid. However studies suggest that the gastric epithelial cells are considerably resistant to acid even at high concentrations (Sanders et al.,

1985), due to the “gastric mucosal barrier”. The gastric mucosal barrier is a complex system made up of epithelial, mucus and submucosal components. The mucus layer is an organized thick layer produced by mucus-neck cells within the epithelium (Clamp &

Ene, 1989). Mucus is comprised of mucin proteins that are heavily glycosylated. Mucus is relatively resistant to proteolysis, it retains water in an unstirred layer, it excludes large molecules and the carbohydrates contents of the glycoproteins mirror that at the surface of the epithelial cell, creating a decoy for microbes (Allen & Garner, 1980).

Additionally, diffusion of hydrophilic molecules is considerably lower in mucus, which causes delayed diffusion of a variety of damaging chemicals, including gastric acid or enzymes, to the epithelial surface (Engel et al., 1995; Allen & Flemström, 2005; McColl,

2012). In addition to the mucus layer, gastric epithelial cells secrete bicarbonate ions

into the lumen, which serves to maintain a neutral pH along the epithelial plasma membrane (Phillipson et al., 2002; Baumgartner & Montrose, 2004). Bicarbonate secreted from the epithelium has been observed to be concentrated within surface mucus, which creates an environment closer to neutrality than the acidic nature of the luminal gastric contents (Wallace, 2008). The pre-epithelial barrier of bicarbonate and mucus is the first-line of defense against the noxious contents of the stomach lumen.

Interestingly, interruption of the mucus-bicarbonate layer does not generally cause or result in epithelial damage (Wallace, 1989). Another component of the gastric mucosal barrier is mucosal hydrophobicity. Epithelial cell membranes lining the stomach wall contain phospholipids, which repel water-soluble luminal contents such as hydrogen. As a result, hydrophobic interactions prevent acid and pepsin back-diffusion. The submucosal component of the barrier is the high rate of mucosal blood flow supplied by a dense network of submucosal capillaries (Bonagura & Kirk, 1995). The mucosal blood flow supplies oxygen and nutrients to the epithelial cells to meet the high metabolic demand for production of gastric secretory products and cell renewal (Bonagura & Kirk,

1995). Gastric mucosal blood flow is also vital in the disposal of intracellular hydrogen ions and the sustainment of chloride transport (Slomiany et al., 1987).

The component of the gastric mucosal barrier we are most interested in is the gastric epithelium. The epithelial lining of the stomach provides a protective barrier that is essential to maintain tissue integrity. The gastric epithelium is renewed every 2-4 days in humans and every 1-3 days in mice , allowing the epithelial barrier to be resilient to injury (Wright, 1984; Matsuo et al., 2017). The epithelium continuously, and quickly, repairs when cells are injured in a process known as epithelial restitution. Epithelial

restitution, which in the initiating step in repair, involves migrating epithelial cells which extend large lamellipodia over the damaged mucosa to quickly (<1 hour) cover small erosions and re-establish an intact epithelium. Restitution also allows for the rapid replacement of damaged cells without compromising the epithelial barrier, thereby preventing further damage to the mucosa.

1.2 Disruption of barrier function

Since the gastric mucosa is continually exposed to a variety of luminal factors which can cause local epithelial damage, it is essential for the epithelium to have the ability to quickly repair. Despite its robust and multi-component nature, the gastric mucosal barrier can be breached. Exposure to chemicals or medications such non-steroidal anti- inflammatory drugs (NSAIDS), physical insults, local infections by pathogens such as

Helicobacter pylori, and a variety of systemic diseases can lead to disruption of the barrier (Tarnawski, 2005; Kusters et al., 2006). Additionally stress, which can be an integral part of illness and trauma, can decrease mucosal blood flow and thereby compromise the integrity of the mucosal barrier (Zhan et al., 2002). Reduced mucosal blood flow suppresses production of mucus and limits the ability to remove any back- diffusing hydrogen ions. As a consequence, significant stress is associated with gastric mucosal erosions. Damage to the gastric mucosa can be mild and readily repaired, or extensive and result in ulcers. Additional insults to the gastric tissue can become persistent or recurring ulcers (Kusters et al., 2006). Peptic ulcer disease (PUD) affects more than 6 million people each year in the United States and continues to be a burden in medical care (Feinstein et al., 2010). Statistics from the NIH report that PUD cost the

U.S. healthcare system 1.9 billion dollars in 2004. Despite decreases in PUD incidence over the past decades, PUD still remains the lead diagnosis on 37% of hospital discharges (Everhart & Ruhl, 2009). Furthermore, new research suggests that proton pump inhibitors (PPIs), the most widely used treatment regimens for PUD, may increase the risk of dementia (Gomm et al., 2016) and chronic kidney disease (Moledina &

Perazella, 2016; Xie et al., 2016). These statistics emphasize the current need to define the mechanisms of gastric repair and identify possible therapeutics. The development of strategies to reduce ulcer incidence or accelerate the healing process represents an important goal for gastric research. The critical initial task following disruption of the gastric mucosal barrier is to cover the denuded area and re-establish the intrinsic barrier. When damage is extensive, the healing process is divided into four distinct phases: (1) restitution, (2) immune activation, (3) proliferation, and (4) differentiation

(Grazul-Bilska et al., 2003; Aihara et al., 2017). This rapid restoration of a continuous epithelium is accomplished by restitution. However, while the injury area is protected, it may require more work to become physiologically functional. To become functional, the subsequent three steps of healing are required. The healing process involves the immune system, particularly macrophages for removal of cellular debris, cellular proliferation, and differentiation of new cells to restore the normal cellular architecture and function. This series of events repairs extensive damage, such as ulceration. In the case of small localized damage that is restricted to the surface epithelium, no immune cell activation, proliferation or differentiation is needed to restore a normal tissue function. In this setting, only restitution is required for proper repair. Restitution occurs

rapidly and coordinates the removal of damaged cells and the migration of healthy epithelial cells into the breach (Figure 1.3).

Physiologically, in Injury Lumen both extensive and local damage, Basement membrane efficient restitution is Migration &

Dead cell important to limit fluid exfoliate and electrolyte loss as well as to prevent Cover denuded luminal antigens and area

bacteria from Restitution accessing the host Restored barrier tissue. It has been speculated that Figure 1.3 Epithelial Repair Process. Following cell injury, dead or damaged cells are expelled into restitution occurs in the gastric lumen and neighboring viable cells migrate to cover the denuded area caused by the departure of the damaged cells. This results in a restored epithelial continuity and barrier function. response to small localized damage, but deficient repair or continual injury can lead to larger more extensive damage. As a result, it is important to understand the mechanisms which mediate gastric restitution.

1.3 Gastric epithelial restitution

After small focal damage, epithelial restitution can be incredibly rapid. In vivo, microlesions quickly repair within minutes in the mammalian stomach (Xue et al., 2010;

Xue et al., 2011). In the case of small damage, restitution is the only event required for repair. Restitution relies upon the rapid response and ability of viable cells to fill in denuded areas and provide the force to expel damaged cells away from the epithelial barrier. Restitution is an intrinsic function of GI epithelial cells that can occur in the absence of systemic signals. Restitution has been demonstrated in vitro in cell lines

(Svanes et al., 1982; Svanes et al., 1983) and recently in gastrointestinal organoids

(Schumacher et al., 2015a; Aihara et al., 2018). Migration of cells to the site of damage generally involves the following: 1) the orderly dissociation of cell–cell contacts, 2) disruption of cell–substratum contacts (focal adhesions) to disconnect viable cells from their site of origin, 3) assembly of actin stress fibers, 4) the formation of leading edge lamellipodia to promote cell movement and 5) motogenic signal production, which prevent cell-substratum disruption-driven anoikis, a form of apoptosis (Taupin &

Podolsky, 2003; Aihara et al., 2017). All these events are necessary to promote cell migration and prevent cell death in neighboring viable cells (Svanes et al., 1982).

Various molecular signals are involved in each step of this process. Restitution has been shown in vitro and in vivo to involve a variety of factors that can regulate the speed of the process including: luminal acid, calcium, epithelial cell microfilaments, prostaglandins, secreted growth factors such as epidermal growth factor (EGF), and trefoil factors (TFFs) (Svanes et al., 1982; Svanes et al., 1983; Xue et al., 2010; Xue et al., 2011; Szabo, 2014).

1.4 Trefoil factor family peptides

One important factor that has been shown to affect gastric epithelial restitution is the mucus-neck cell secreted peptide, trefoil factor (TFF). TFF peptides are a family of secretory molecules that are responsible for mediating several physiological roles that maintain and restore GI mucosal homeostasis. There are three members in the mammalian TFF gene family: TFF1, TFF2, and TFF3 (Aihara et al., 2017). TFF peptides are widely expressed in various tissues; however they are detected at high levels in the

GI tract especially in the mucosal layer. In the stomach, TFF1 and TFF2 are predominantly expressed, though there are negligible traces of expression of these peptides further down the GI tract. TFF1 is predominantly expressed within the gastric foveolar cells as well as the surface epithelial cells throughout, however studies suggest that TFF1 primarily acts as a tumor suppressor within the gastric epithelium (Aihara et al., 2017). In both rodents and humans, TFF2 is expressed in similar cell locations in gastric epithelial cells of the corpus and antrum regions and gastric mucous neck cells

(Jørgensen et al., 1982; Hanby et al., 1993; Lefebvre et al., 1993). TFF2 has been implicated in acting as a mucosal protective peptide as well as regulating/facilitating repair (Playford et al., 1995; Babyatsky et al., 1996; McKenzie et al., 1997; Xue et al.,

2011). All TFFs have been shown to be motogens (Jørgensen et al., 1982; Masiakowski et al., 1982; Suemori et al., 1991; Xue et al., 2011), substances that stimulate cell motility, having the ability to promote migration of viable cells to cover the denuded area without proliferation (Tomasetto et al., 2000). While TFF2 is known to be motogenic, the underlying mechanism of TFF2-driven restitution remains largely unknown. Numerous factors have been identified in vitro and in vivo to regulate cell migration and epithelial

restitution, including calcium, epidermal growth factor (EGF), sodium hydrogen exchanger isoform 2, and TFF peptides (Tomasetto et al., 2000; Kjellev et al., 2006; Ota et al., 2006; Albert et al., 2010; Xue et al., 2010).

In the context of gastric ulcer healing, TFF2 is upregulated and is localized in cells that also express the gastric chief cell marker intrinsic factor. The complex process of gastric ulcer healing requires cellular proliferation, remodeling of both mucosa and blood flow in the tissue, as well as cell migration to cover the denuded tissue (Tarnawski et al., 1991; Okabe & Amagase, 2005). During ulcer healing, a reparative lineage emerges that is characterized by cells that are dual positive for mucous neck cell makers (MUC6, GSII and TFF2) and gastric chief cells (intrinsic factor). This gastric lineage is referred to as spasmolytic polypeptide expressing metaplasia (SPEM)

(Nomura et al., 2004; Ogawa et al., 2006; Nozaki et al., 2008). SPEM also arises following loss of gastric parietal cells in a variety of models including L635, DMP777, high dose tamoxifen administration and H. pylori infection. In general, the native gastric corpus mucosa does not express (or has low expression) many SPEM markers, such as Clusterin, HE4 (Wfdc2), Dmbt1, Cxcl17, Cenpk, Top2a, Ube2c, and others (Weis et al., 2013; Weis et al., 2014; Engevik et al., 2016). However, following loss of gastric parietal cells or injury there is a dramatic increase in gene expression of the above- mentioned genes (Weis et al., 2014; Engevik et al., 2016). During gastric ulcer repair,

SPEM emerges in the base of the ulcer margin (Kikuchi et al., 2010; Engevik et al.,

2016). The absence of SPEM in aged mice following injury results in improper mucosal healing suggesting that SPEM is likely necessary for normal gastric epithelial regeneration (Engevik et al., 2016).

Numerous studies have demonstrated a mucosal protective effect of TFFs in the

GI tract. In mutant mice that lack any one of the TFF proteins, there is an increased susceptibility to epithelial damage in injury models (Mashimo et al., 1996; Farrell et al.,

2002). In complementary studies, exogenous TFFs have been demonstrated to oppose the generation of injury by numerous agents. Both approaches suggest that TFFs contribute to mucosal defenses in the normal tissue. Exogenous TFF2 has been shown to inhibit gastric injury induced by nonsteroidal anti-inflammatory drugs (NSAIDs) or ethanol (Babyatsky et al., 1996; McKenzie et al., 1997; McKenzie et al., 2000). TFF2

KO show an increase susceptibility to NSAID injury in the stomach (Farrell et al., 2002).

It should be noted that, at experimentally applied concentrations, TFF peptides have been shown to be promiscuous as similar physiological effects can be elicited by any one of the TFF isoforms when added as exogenous peptides (Kjellev et al., 2006). Also of note, TFF peptide has been detected in both luminal fluid and serum of the GI tract

(Dubeykovskaya et al., 2009). The general assumed route of TFF2 action is via luminal secretion from the GI epithelial cells and study results suggest that systemic TFF peptides are able to be secreted into the gastric lumen parallel with mucus secretion from mucous cells (Aikou et al., 2011). Regardless of the promiscuity of TFF peptides,

TFF2 has at least one known receptor in the stomach, C-X-C chemokine receptor 4

(CXCR4). In epithelial cells, TFF2 peptide is observed to directly activate CXCR4. It has been observed in gastric cancer-derived epithelial cell lines that CXCR4 responds to

TFF2 treatment with a robust phosphorylation of ERK1/2 (Rosenstiel et al., 2007).

Furthermore, in vivo and in vitro CXCR4 activation has been noted to require high

concentrations of recombinant TFF which is consistent with reports of high TFF levels under normal physiological conditions (Rosenstiel et al., 2007; Buda et al., 2012).

Due to the complexity of examining cell migration in vivo, most migration studies have utilized in vitro cancer cell lines which have provided the majority of information about TFF signaling and motogenic activity (Aihara et al., 2017). In cancer-lines, scratch assays are the most common method for examining restitution. However, gastric cell lines require >24 hours for epithelial restitution (Zheng et al., 2013; Wang et al., 2018).

Even though application of TFF in the gastric cancer cell lines SGC7901 and AGS does improve restitution compared to controls, full closure of the wound area is not observed at 24 hrs (Yu et al., 2010; Zheng et al., 2013). This repair phenotype does not mirror restitution in vivo, which occurs rapidly (Xue et al., 2010; Xue et al., 2011; Aihara et al.,

2013).

1.5 Epithelial Growth Factors in gastric restitution

Several studies from in vitro cancer cell lines have also pointed to the role of the

ErbB pathway in TFF signaling. Epidermal Growth Factor Receptor (EGFR), also known as ErbB1, is present on the gastric epithelial cells (Mori et al., 1987; Ménard & Pothier,

1991; Chen et al., 2001). EGFR recognizes the ligand epidermal growth factor (EGF) and stimulates migration, proliferation and survival within cells. In healing gut mucosa, it has been noted that EGF receptor (EGFR) is highly expressed apical to the lumen

(Hansson et al., 1990). In rabbit and porcine models of experimental ulceration, treatment of ulceration by EGF-containing chitosan hydrogel resulted in enhanced ulcer healing, potentially due to the prolonged release of EGF and a concomitant sustained

activation of EGFR for an extended period of time (Maeng et al., 2014). In human gastric epithelial monolayers, addition of EGF, which subsequently triggered phosphorylation and activation of EGFR, greatly accelerated wound healing (Tétreault et al., 2008).

Outside of the gastrointestinal tract, evidence points to the potential of TFF2 acting through the TFF receptor CXCR4, a G-coupled protein receptor (GCPR), to transactivate EGFR (Kinoshita et al., 2000; Rodrigues et al., 2003). In addition to ligand binding by the EGFR receptor itself, activation of EGFR upon activation of other GPCRs has been established as mechanism of activation; this is known as EGFR transactivation. GPCR mediated EGFR transactivation occurs through an initial heterotrimeric G protein dissociation, activation of ligand-specific intermediates including non-receptor tyrosine kinase, followed by a metalloprotease activation

(Forrester et al., 2016). There is evidence that TFF2 peptides signal through ErbB pathways, of which EGFR is a member (Emami et al., 2004). Additionally, apart from

TFF, in healing gut mucosa, studies have shown the presence of stimulated EGFR and

MAPK (ERK1/2) (Emami et al., 2004). However, it is unknown whether this signaling is due to direct interaction of TFF2 with EGFR/ErbB or if another intermediate EGF family ligand is involved. In human gastric cancer cell lines, it has been observed that incubation with recombinant TFF2 leads to EGFR phosphorylation and MAPKs activation (Taupin et al., 1999). TFF2 KO mice exhibit delayed gastric repair, however loss of TFF2 results in no change in two major gastric EGFR ligands (Farrell et al.,

2002). These results suggest that TFF2 may be required for full action of EGF signaling in the stomach and provides evidence for the role of EGFR in gastric restitution.

1.6 Sodium hydrogen exchangers in gastric restitution

Sodium hydrogen exchangers (NHE) have been demonstrated as key modulators of

Na+ absorption, internal pH [pHi], cell volume and gastric and intestinal wound repair

(Furukawa & Okabe, 1997; Furukawa et al., 1999; Yanaka et al., 2002; Moeser et al.,

2008). Work from our lab has shown that TFF2 requires the sodium hydrogen exchanger isoform 2 (NHE2) for proper repair following damage (Xue et al., 2011). In vivo, NHE2 KO and NHE2 inhibitor treated WT mice have delayed restitution and repair of microscopic lesions, suggesting an important role during repair (Xue et al., 2011).

NHE2 has been implicated in vivo as an important downstream effector of TFF2 signaling as addition of recombinant TFF2 is inhibited by NHE2 inhibitor in TFF2 KO mice. This evidence suggests NHE2 acts as a mediator of TFF proteins action to promote gastric epithelial restitution (Xue et al., 2011). Additionally, EGF contribution to restitution has been found to be mediated in part by stimulation of NHE in gastric epithelial cells (Vexler et al., 1996; Yanaka et al., 2002) and EGF is involved in acute regulation of NHE activity (Iwatsubo et al., 1989; Ghishan et al., 1992; Furukawa &

Okabe, 1997). In rats in vivo and in vitro studies, EGF treatment increased NHE2 activity and mRNA abundance by nearly twofold (Xu et al., 2001). The ERK pathway has also been shown to a critical component of NHE activation. In intestinal Caco-

2BBe1 cells, activation of ERK and PKC stimulates NHE2 expression (Muthusamy et al., 2011). Lysophosphatidic acid 5 (LPA5)-dependent activation of NHE3 in Caco-2BBe cells requires the mitogen-activated protein kinase MEK and ERK; an effect mediated by transactivation of EGFR (Yoo et al., 2011). These studies indicate a potential link

between TFF, EGFR and NHE2. Our work herein describes the first study to connect these three components and identify the signaling cascade associated with repair.

1.7 Role of calcium in repair

Calcium (Ca2+) has been known as an effector of gastric wound repair since

1985 (Critchlow et al., 1985; Takeuchi et al., 1985; Cheng et al., 2001), but the mechanism by which Ca2+ mobilization affects restitution has yet to be elucidated. As a ubiquitous second messenger, Ca2+ influences numerous cellular processes, including mucus secretion and cell migration in gastric epithelial cells (Belkacemi et al., 2005;

Schreiber, 2005). It has been observed in cultured rabbit gastric epithelial cells that intracellular Ca2+ is present in significantly higher amounts in migrating cells at the edge of a scratch wound 2 hours following damage (Ranta-Knuuttila et al., 2002).

Furthermore, in these cells treatment with verapamil (a calcium channel blocker), calphostin-C (PKC inhibitor) and calmidazolium (calcium/calmodulin complex inhibitor) significantly inhibit cell migration speed observed at 24 hours following monolayer wounding (Ranta-Knuuttila et al., 2002). Intracellular Ca2+ is demonstrated to be essential for actomyosin contractile forces (Strohmeier & Bereiter-Hahn, 1984; Citi &

Kendrick‐Jones, 1987; Rees et al., 1989), regulation of actin cytoskeleton structure and dynamics (Hartwig & Yin, 1988; Condeelis, 1993), maintenance of adherens and tight junctions (Gonzalez-Mariscal et al., 1990; Marchiando et al., 2011), and cell-substratum adhesion assembly or disassembly (Crowley & Horwitz, 1995; Sjaastad & Nelson,

1997). Ca2+ has been shown to be especially important in regulation of myosin light- chain kinase, which is essential for myosin activation to increase cell contractility in

migrating cells (Ridley et al., 2003; Aihara et al., 2018). In vitro, extracellular Ca2+ has been demonstrated as an essential factor in regulation of actin dynamics, potentially in the maintenance of cellular tightness and motility (Aihara et al., 2018).

In vivo in mice, both intracellular and extracellular calcium have been shown to be essential for proper gastric wound repair (Aihara et al., 2013). Calcium signaling has been reported to be activated by TFF2 via CXCR4 (Dubeykovskaya et al., 2009), however there is no direct evidence that calcium mobilization is required for TFF mediated repair. Furthermore, activation of CXCR4 in the Caco-2 cell line was found to stimulate the release of intracellular Ca2+ and enhance intestinal epithelial restitution through reorganization of the actin cytoskeleton via the calcium-dependent focal adhesion kinase Pyk2 (Agle et al., 2010). It has also been reported that gastric epithelial damage is associated with intracellular and extracellular Ca2+ mobilization in vivo and this influx of calcium is required to mediate tissue repair (Aihara et al., 2013).

The key role of Ca2+ in repair has also been confirmed by the use of inhibitiors.

Addition of PLC inhibitor, IP3 receptor antagonist, or COX inhibitor slowed gastric repair while also blocking calcium mobilization stimulated in the restitutive epithelial cells

(Aihara et al., 2013). These drug effects are suggested to be mediated by the inhibition of Gq protein activation (Aihara & Montrose, 2014). Chelation of intracellular calcium by

BAPTA/AM has been shown to inhibit the increase of luminal calcium in vivo (Aihara et al., 2013), this suggests that the increase of luminal calcium (which benefits epithelial repair) is dependent on intracellular calcium increase. Activation of phospholipase C

(PLC) is a known initiator of calcium-dependent signaling. Once the PLC pathway is activated, PLC cleaves the lipid phosphatidylinositol to release inositol triphosphate

(IP3) and diacylglycerol (DAG), metabolites that can stimulate calcium release from intracellular stores and activate protein kinase C (PKC). PKC is also a conventional target of calcium-dependent regulation. Both PLC and PKC have been shown in multiple cell types to stimulate epithelial cell migration (Ranta-Knuuttila et al., 2002; Rao et al., 2007; Saidak et al., 2009). In human gastric cancer (AGS) cells, PKC activation and calcium mobilization have been shown to stimulate repair from aspirin and deoxycholate induced damage (Redlak et al., 2007, 2008). The calcium-dependent PKC alpha isoform and PLC-beta isoforms are known to be localized in gastric surface cells

(Miller & Henagan, 1979; McGarrity et al., 1996), however the gastric defensive role of

PKC and PLC has yet to be evaluated.

In rats, gastric damage by addition of taurocholate, 1 M NaCl, or 50% ethanol elicited increased gastric luminal Ca2+ as an adaptive response to prevent further damage (Takeuchi et al., 1985; Koo, 1994; Takeuchi et al., 1999). In frog gastric mucosa, extracellular Ca2+ was required for restitution following hyperosmotic injury

(Critchlow et al.). In mice in vivo, intracellular and extracellular calcium have been demonstrated to be necessary for proper gastric wound repair (Aihara et al., 2013).

TFF2 is known to activate Ca2+ signaling via the CXCR4 chemokine receptor

(Dubeykovskaya et al., 2009), however there is no direct evidence that Ca2+ mobilization is required for TFF mediated repair. While these studies point to the overarching role of endogenous Ca2+ in gastric epithelial repair, little is known about the early signaling events driving Ca2+ mobilization.

1.8 Genetically encoded calcium indicators

While intracellular loading of conventional acetoxy methylester calcium-sensitive fluorescent probes, such as fura-2 or fluo-4, have been utilized in in vitro, for unknown reasons these probes fail to load effectively in vivo in normal functioning gastric cells

(Aihara et al., 2013). Due to past limitation of calcium sensors and difficulty in monitoring intracellular calcium or wound repair in real time, the mechanistic basis of calcium mobilization in healthy tissue has been largely unexplored.

Recently fluorescent protein-based indicators have been used as an invaluable tool for live imaging of living cells and organisms. The use of genetically encoded Ca2+ indicators (GECIs) is a particularly promising approach since cells display a transient increase in intracellular Ca2+ concentration when they receive specific intercellular signals. GECIs, including yellow cameleons (YCs) and GCaMPs/pericams, represent a highly advanced class of indicators (Miyawaki et al., 1997; Nagai et al., 2001; Nakai et al., 2001). Improvements in the performance of GECIs pH sensitivity, wavelength for absorption and emission, dynamic range and improved Ca2+ affinity have resulted in brighter indicators with superior photo-stability and expanded dynamic range; thereby improving the calcium sensitivity. The YC system uses a fluorescence resonance energy transfer (FRET) mechanism to report on Ca2+ signals. In this system, a cyan fluorescent protein (CFP) is linked to a yellow fluorescent protein (YFP) by a calcium- binding domain, calmodulin and M13 peptide which can bind to the calcium-bound form of calmodulin (Arai & Nagai, 2013). One recent improvement for the YC system was the modification of the linker peptide between CaM and the M13 peptide (5 amino acids longer; Gly-Gly-Gly-Gly-Ser). The resulting modification resulted in a higher Ca2+ affinity

(Kd = 15 nM) compared with those with a shorter linker (Horikawa et al., 2010). This YC protein was termed 'YC Nano15' and currently this indicator has the highest affinity of any GECI reported so far (Horikawa et al., 2010).

A potential caveat of GECI use is that the strong Ca2+ chelating effect of YC

Nano might affect endogenous Ca2+ homeostasis. This phenomenon has been observed for other molecules or ions, such as cyclic nucleotides and NO, when using loaded indicators. However, this is not the case for Ca2+. Similar to H+, free cytosolic

Ca2+ is maintained dynamically through the balancing action of Ca2+ buffers (i.e., Ca2+ binding proteins), that are abundant within the cell (Cheng & Lederer, 2008; Nagai et al.,

2014). As a result, GECIs offer significant advantages over synthetic Ca2+ dyes in their targetability and reliability for long-term imaging. GECIs, used in collaboration with high resolution microscopy represent an innovative advanced system to monitor calcium mobilization during epithelial repair. Moreover, the use of gastric organoids containing

GECI allow the ability to bypass systemic effects of inhibitors or agonists while also reflecting expression similar to native tissue. To the best of our knowledge, our work with YC Nano15 gastric organoids represents the first documentation of Ca2+ mobilization in response to epithelial damage and TFF2 in any organoid model.

1.9 Gastric cancer lines vs. gastric organoids

Much remains unclear due to complexities of in vivo work and the inconsistencies of in vitro immortalized or cancer-derived cell lines. Cancer-derived cell lines are known to differ from native tissue, especially in the areas of cell growth, migration, and differentiation (Xu et al., 2017). For example, the histone demethylase KDM5C and p53

that promotes cancer cell growth is enriched in cancer-cell lines compared to normal gastric tissue (Xu et al., 2017). Cancer-derived cell lines also exhibit lower expression of

Calpain-8, calpain-9, ERp29, miR-30b, miR-370, miR-377, miR-194-4p and tristetraprolin (TTP) compared to normal gastric mucosa (Zheng et al., 2013; Zhu et al.,

2014; Deng et al., 2016; Peng et al., 2016; Wu et al., 2017; Bo et al., 2018; Wei et al.,

2018). Additionally, gastric cancer lines exhibit higher expression of RASSF8, SOX20T,

GCRK1, and AK069174 compared to normal gastric tissue (Bo et al., 2018; Lin et al.,

2018; Wei et al., 2018; Zhang et al., 2018). In terms of signaling cascades, IL-26 and

STAT3 signaling is activated in human gastric cancer cell lines compared to non-cancer controls (You et al., 2013). Not only do cancer-cell lines differ from normal gastric tissue, gastric cancer-derived lines also differ from their original cancer samples. It has been shown that genetic alterations occur within gastric cancer lines compared to their original gastric cancer tissue, particularly in the genes microsatellite instability (MSI), loss of heterozygosity (LOH) and p53 (Bae et al., 2000). While not an ideal model, due to the altered regulation of proliferation and cell migration (invasion), cancer cells have provided the bulk of information about the signaling cascades involved in restitution that relate to cell migration.

In contrast to cancer-derived cell lines which differ from native cancer tissue and normal gastric tissue, gastric organoids appear to resemble normal gastric tissue.

Engevik et al. recently demonstrated by RNAseq analysis that mouse fundic gastric organoids grown in culture for 7 days exhibited a similar gene expression pattern to native mouse fundic tissue (Engevik et al., 2016). While current evidence points to the usefulness of gastric organoids to allow for studies that align more with in vivo native

tissue, limitations still exist within this model as gastric organoids do not fully recapitulate native tissue. While gastric organoids have been demonstrated to have similar physiological functions as in vivo, the exact gland architecture and cell arrangement are not present within the organoid model (Schumacher et al., 2015a).

Furthermore while functional parietal cells are present in the organoid, the overall distribution and population is decreased compared to in vivo (Schumacher et al.,

2015a). Of note, the organoid model can be improved by co-culturing gastric organoids with immortalized mesenchymal cells which maintains the number of parietal cells similar to native tissue (Schumacher et al., 2015a; Bertaux-Skeirik et al., 2016). Due to a decreased population of parietal cells, as well as increased mucus cells and metaplastic markers (Engevik et al., 2016), there is concern organoids may represent more metaplastic than native tissue. RNA sequencing data suggest that gastric organoids highly express a number of genes that are identified with SPEM including

TFF2, Gpx2, Clusterin, Dmbt1, Cxcl17, Cenpk, and Top2a among other genes (Engevik et al., 2016) compared to gastric glands. This indicates that gastric organoids may more closely mimic the metaplastic SPEM lineage than the native fundic epithelium.

Transplantation of gastric organoids generated from young mice into the gastric mucosa of aged mice following injury resulted in the emergence of SPEM and normal gastric regeneration in aged mice (Engevik et al., 2016). This provides further evidence that gastric organoids may represent metaplastic cells that are capable of promoting repair.

Since gastric organoids highly express a number of metaplastic genes studies using gastric organoids may not fully recapitulate in vivo epithelial cells. However, currently gastric organoids are the only gastric in vitro primary culture system that can be

extensively used and is not derived from cancer cells or transformed cells making it the most amenable in vitro model available as a simplified representative model of the gastric epithelium.

1.10 Major Contributions

The work herein describes the first use of GECI gastric organoids with 2-photon damage monitoring gastric restitution in real time. We have identified pathways involved in TFF2, EGFR, ERK1/2, NHE2 and Ca2+ signaling during gastric epithelial repair, confirming previous in vivo findings and expanding our current understanding of the repair mechanism. These mechanisms link pathways which previously existed in isolation. This innovative examination of epithelial restitution will likely have a positive impact on our ability to improve therapeutics for promoting gastric repair.

Chapter 2

Assessing permeability, repair and cell death in the gastric organoid system

Chapter 2 Assessing permeability, repair and cell death in the gastric organoid system

2.1 Abstract

Localized injury to the gastrointestinal (GI) epithelium requires a rapid response to prevent further damage known. During restitution, the initial event in repair, the damaged cell is exfoliated into the lumen while neighboring viable cells migrate to cover the damaged area. With the establishment of 3D primary culture of gastric epithelial cells, known as gastric organoids, we use two-photon induced photodamage (PD) to test the hypothesis that gastric organoids can serve as a useful in vitro model to assess gastric restitution with quantitative measurements of repair as well as address the type of cell death associated with PD. Gastric organoids were generated from isolated fundic tissue of wild-type (WT) or TFF2 knockout (KO) mice, or Yellow Cameleon Nano15 (YC

Nano) transgenic mice. In WT and TFF2 KO gastric organoids, permeability was assessed based upon intensity of Lucifer yellow (LY) added in culture medium as an extracellular marker. In unperturbed organoids, LY did not leak into the luminal space of the gastric organoids, confirming integrity of the epithelial barrier. Following PD, LY did not significant increase in the organoid lumen of either WT or TFF2 KO organoids compared to unperturbed state. In EDTA supplemented WT gastric organoids, following

PD, LY significantly increased in the organoid lumen and significantly delayed exfoliation compared to WT. In WT and TFF2 KO gastric organoids, progression of damage and repair was evaluated based upon damaged cell nuclei movement (10

µg/ml Hoechst33342). Following PD, exfoliation in TFF2 KO organoids was significantly delayed compared to WT. In YC Nano gastric organoids, which contain a yellow fluorescent protein that marks the cytosol, the damage area was filled within 10 min

following PD. To assess cell death, caspase 3/7 detection reagent (5µM) was added to

WT organoids, fluorescence was monitored before and following PD. In unperturbed organoids, caspase 3/7 detection reagent was only observed in natural cell shedding.

PD to a single cell elicited caspase-activated apoptosis and resulted in maximum exfoliation of the dead cell by 10 min. Addition of Z-VAD-FMK (pan-caspase inhibitor) confirmed reliability of caspase 3/7 detection reagent as an indicator of caspase- activated apoptosis, as well as demonstrated the necessity of caspase-activation for proper restitution. Gastric organoids mirror restitution response observed previously in vivo studies. Two-photon induced damage results in caspase-activated apoptosis, which mirrors unperturbed natural cell shedding. Damage area and exfoliation are useful indicators of repair within gastric organoids, while LY is useful as an indicator of epithelial barrier integrity.

2.2 Introduction

The gastric epithelium acts as a primary barrier against the noxious contents of the stomach, where the acidic nature of the gastric lumen creates a unique and challenging physiological environment. In response to injury of the epithelial lining, dying cells are immediately expelled into the lumen while viable neighboring cells simultaneously cover the denuded area. This process is known as restitution and occurs within minutes following injury in vivo (Demitrack et al., 2010; Xue et al., 2010; Xue et al., 2011; Aihara et al., 2013). In vitro microlesions caused by two-photon induced photodamage (PD) also elicit rapid restitution in gastric organoids (Schumacher et al.,

2015a; Aihara et al., 2018). The induction of microlesions via two-photon microscopy

has been well established as a model of cell damage in vivo (Nyqvist et al., 2005; Xue et al., 2010; Xue et al., 2011; Aihara et al., 2013) and recently in vitro in the gastric organoid system (Schumacher et al., 2015a; Aihara et al., 2018).

While this damage model has reproducible results, it has not been fully investigated as to the type of cellular death that is induced via two-photon PD. In vitro, the two-photon induced PD model has been primarily examined in gastric organoids derived from transgenic mouse expressing fluorescent proteins (Schumacher et al.,

2015a; Aihara et al., 2018) to address epithelial restitution based upon the closure of the damage area indicated by loss of fluorescence. Gastric organoids have been utilized to study an array of disease states (Schumacher et al., 2015a; Bartfeld, 2016;

Engevik et al., 2016; Engevik et al., 2018b), there are limited studies in quantitatively assessing epithelial repair and its signaling cascade within this system (Aihara et al.,

2018). Additionally, there is limited information regarding cell turnover under normal conditions. In other in vitro epithelial models, cell turnover has been demonstrated to be due to cell death via apoptosis either by caspase-activation, anoikis, or Sphingosine 1

Phosphate signaling (Gudipaty & Rosenblatt, 2017). In corneal epithelium, cell turnover has been shown to rely upon more than one mechanism (ie apoptosis) to remove cells from the epithelial barrier to maintain its function (Ren & Wilson, 1996). In human stomach, caspase3 has been suggested to regulate apoptotic cell death in cell turnover in normal mucosa (Hoshi et al., 1998). Studies involved in gastric epithelial cell turnover primarily focus upon the role of the pathogen, Helicobacter pylori, to induce apoptosis and alter gastric epithelial cell turnover (Moss et al., 1996; Peek et al., 1999; Cover et al., 2003). This study seeks to address approaches that have potential to assess repair

within non-fluorescent organoids as well as determine if the cell death induced by PD is related to what occurs during normal cell turnover in organoids.

2.3 Materials and Methods

2.3.1 Animal husbandry

Experiments used TFF2 wildtype (WT) (C57BL/6 background) or TFF2 knockout

(KO) (C57BL/6 background) mice (Xue et al., 2011), or transgenic mice (C57BL/6 background) expressing the Yellow Cameleon-Nano15 (YC Nano) Ca2+ sensor fluorescent protein (Oshima et al., 2014). Pups were genotyped by genomic PCR as previously described (Schultheis et al., 1998; Bell et al., 1999; Farrell et al., 2002) and used for experimentation at 2-3 months of age. Animals were given standard rodent chow diet and water, both ad libitum. All animal procedures were approved by the

Institutional Animal Care and Use Committee of the University of Cincinnati.

2.3.2 Mouse-derived corpus organoid culture

Gastric organoids were generated from mouse gastric corpus as described

(Mahe et al., 2013; Schumacher et al., 2015a; Engevik et al., 2018b). Isolated gastric epithelium from the corpus was cultured in Matrigel diluted 1:1 in Dulbecco’s

Phosphate-Buffered Saline (DPBS) without Ca2+ and Mg2+ in 8-well or 2-well Lab-Tek chamber with coverglass (Thermo Scientific) to grow gastric organoids. Gastric organoids were cultured in a 5% CO2 incubator at 37°C for 3-4 days prior to experiments.

2.3.3 Induction of two-photon laser-induced photodamage

Experiments were performed in organoid culture medium under 5% CO2/37 °C conditions in a microscope incubation chamber (PeCon, Erbach, Germany) on an inverted confocal microscope (Zeiss LSM 510 NLO) and imaged with a C-Achroplan

NIR 40x objective lens. In some experiments using WT gastric organoids, gastric organoids were pre-incubated for 30 min with the DNA stain Hoechst 33342 (10 μg/ml,

Invitrogen) to visualize cellular nuclei. In experiments intended for analysis of damage area in YC Nano gastric organoids, images of YFP (excitation 514 nm, emission 535–

590 nm) in the gastric organoid were collected simultaneously with transmitted light and a confocal reflectance image (reflecting 730 nm light to show cell/tissue structure). In

WT gastric organoids, images of Hoechst 33342 (titanium-sapphire laser [Ti-Sa] excitation 730 nm, emission 435–485 nm) were collected simultaneously with transmitted light and confocal reflectance images. In all photodamage experiments, after collecting a set of control images, a small rectangle region (≈5 µm2) of a single cell was repetitively scanned at high Ti-Sa laser power (730 or 840 nm: 630 mW average) for 500 iterations (requiring ≈3 s).

Experiments examined gastric organoids embedded in Matrigel, located approximately 50-300 μm from the cover glass. In experiments testing permeability,

EDTA (2mM, Fluka) was applied to medium and used immediately. In a subset of experiments, Lucifer Yellow (20 µM, Molecular Probes) (Argon laser excitation 458 nm, emission 500-550 nm) was added to the medium of WT gastric organoids. In separate experiments, CellEventTM Caspase 3/7 Green Detection Reagent (5µM, ThermoFisher)

(Argon laser excitation 488 nm, emission 500-550 nm) was added to the media of WT and incubated for 30 min prior to experiments.

Damage-repair cycle was measured independently once per gastric organoid, and outcomes from at least 4 different gastric organoids (derived from at least 3 animals), were compiled for each experimental protocol.

2.3.4 Microinjection

To test maintenance of gastric organoid barrier integrity, Lucifer yellow (20 mM stock) was microinjected into WT gastric organoids using methods previously described

(Engevik et al., 2018b). Gastric organoids (~400-500µm diameter) were injected with

2.3 nL Lucifer yellow for an estimated final concentration of 1mM Lucifer yellow .

2.3.5 Image analysis

All parameters were quantified from the time course of images as described (Xue et al., 2010; Xue et al., 2011; Aihara et al., 2013; Aihara et al., 2014) using Image J and/or

Metamorph software (ver. 6.3, Molecular Devices, Downington, PA, USA). In experiments using Lucifer yellow as an extracellular marker, two separate measurement approaches were used to measure Lucifer yellow intensity in the luminal domain of the gastric organoid. Both approaches compared Lucifer Yellow fluorescence in different physical locations moving away from the damage site. Both approaches normalized results to the intensity of Lucifer yellow in the fluid bathing the organoid (designated at

100%). In the first approach, a line scan (50 µm length and 20 µm width) was positioned perpendicular to the epithelial layer (position 0 µm), extending from the extracellular

space (-20 µm) across the gastric organoid cells and into the lumen of the gastric organoid (up to +30 µm). The line scan intersected the epithelial layer 5-10 µm away from the damage site, with measurements taken on both sides of the damaged area

(see Figure 2.2A). In the second approach, a line scan was positioned entirely in the lumen and roughly parallel to the epithelial layer, with intensity measured as a function of distance away from the damage site, starting at 5-10 µm away from the site of damage (see Figure 2.2D).

The damaged area was measured as the region of cellular loss of YFP fluorescence in YC Nano gastric organoids. In each experiment of YC Nano gastric organoids, we determined the time point displaying maximal damage area and estimated rates of epithelial restitution starting from this time with a single exponential curve fit to the changing size of damaged area over time (Xue et al., 2010; Aihara et al., 2018). Best fit values of the rate constant were used as estimates of the rate of repair (units of min−1).

In WT and TFF2 KO gastric organoids, movement of the damaged cell nuclei was tracked and reported as the maximal distance from the original position attained over the 10 min following photodamage (exfoliation, µm). 10 min was selected as the cut off point for exfoliation because maximal distance occurred at this time point in WT organoids.

In experiments using Caspase 3/7 Detection Reagent, the region of interest (5 µm2) was traced around the damaged cell nuclei (determined by Hoechst33342 stain) to distinguish fluorescence in background corrected images over time.

2.3.6 Statistical analysis

All values were reported from representative experiments as the mean ± standard

error of the mean (SEM) from multiple experiments. Statistical significance was

determined using unpaired Student’s T-test, or one-way ANOVA with Dunnett’s multiple

comparison post-hoc test. A p value of <0.05 was considered significant.

2.4 Results

2.4.1 Assessing organoid permeability using Lucifer yellow

We have observed that microinjection of Lucifer yellow (LY) into gastric

organoids is retained within the lumen for over 3 days (Figure 2.1), suggesting that the

barrier integrity is maintained within the gastric organoid system and sufficient to

severely restrict

transepithelial flux of

this molecule with a

Molecular Weight of

444 g/mol. To assess

the barrier integrity

Figure 2.1 Microinjection of Lucifer yellow into a gastric organoid. Stereoscopic images present in the during Lucifer yellow injection in brightfield (upper panel) or fluorescence (lower panel). Images taken before, during, and immediately, 1 day, or 3 days following microinjection of 2.3 nL LY. organoid system Engevik, K et al. Host-Pathogen Interactions 2018. more acutely, LY was

added to the media as an extracellular marker less than an hour before

experimentation. LY intensity, in the extracellular space and gastric organoid lumen,

was monitored in a timecourse before, and following photodamage (PD). As seen in

Figure 2.2A, in WT control gastric organoids under normal conditions LY was maintained in the extracellular domain with no observable “leakage” into the gastric epithelium or the organoid lumen. Further measurements were made in attempt to identify if the local area around the damaged cell had enhanced leakage of LY.

In our first attempt (Figure 2.2A in images before PD), a line scan was positioned perpendicular to the epithelial layer (position 0 µm), extending from the extracellular space (-20 µm) across the cells into the lumen of the gastric organoid (up to +30 µm). This line scan was positioned to be within 5-10 µm distance from the damage site. Following PD, while the area of the damage cell retained some LY, no additional LY entered into the cell compared to the time point before PD (Figure 2.2B).

To confirm that barrier function was necessary to maintain LY outside of the organoid, the extracellular calcium chelator EDTA (2 mM) was added to the media to weaken the tight junctions (Tomita et al., 1996) (Figure 2.2A). As seen in Figure 2.2A and 2.2C, using the same measurement scheme, EDTA caused an increase of LY within the lumen of the organoid.

As an additional approach to measure LY as an assessment of localized organoid permeability, a line scan was positioned entirely in the lumen and roughly parallel to the epithelial layer (see Methods) (Figure 2.2D in image PD), with intensity measured as a function of distance away from the damage site (Figure 2.2E). At 5 µm away from the damage site, prior to PD, EDTA supplemented gastric organoids demonstrated a significant increase in LY intensity within the organoid lumen compared to WT control (*p<0.05) (Figure 2.2F). At 10 min following PD, WT control did not have a significant increase of LY intensity within the organoid lumen. However, at 10 min

following PD in EDTA supplemented gastric organoids showed a significant increase in

LY intensity in the lumen compared to both WT before PD and EDTA before PD (Figure

2.2F).

This data suggests that the organoid system maintains barrier integrity in a similar manner to what has previously been observed in gastric epithelial cells in vivo

(Demitrack et al., 2010;

Xue et al., 2010; Xue et

Figure 2.2 Approaches to assess organoid permeability using Lucifer yellow. A) Representative images over time course comparing control (upper panel) and 2mM EDTA supplemented (lower panel) WT gastric organoids before and following PD. 2mM EDTA was supplemented to WT gastric organoids 5 min prior to PD. B) Representative measurement from 3 experiments of Lucifer yellow intensity (100%) from line scan across the extracellular space and organoid lumen shown in 4A control gastric organoids (upper panel), results compare before PD, 10 min and 20 min after PD; position zero indicates the epithelium. C) Representative measurement from 3 experiments of Lucifer yellow intensity (100%) from line scan across the extracellular space and organoid lumen shown in 4A EDTA supplemented gastric organoids (lower panel), results compare before PD, 10 min and 20 min after PD; position zero indicates the epithelium. D) Representative images over time course in WT gastric organoids before and following PD. E) Measurement of Lucifer yellow intensity (ratio) along arrow shown in 4D, results compare before and after (10 min) PD in WT (black, n=3) and EDTA supplemented WT gastric organoids (orange, n=3). F) Comparison of Lucifer yellow intensities (ratio) shown in 4E at 5 µm from the damage area (both n=3), *p<0.05 vs WT 0 min; #p<0.05 vs EDTA 0 min.

al., 2011), with no evidence of localized permeability changes that are detectable under normal conditions.

2.4.2 Assessing repair in various organoid models

Our lab has previously used PD in vivo and in vitro as a method to target individual gastric cells (Xue et al., 2010; Xue et al., 2011; Mahe et al., 2013) and monitor restitution by measuring cell exfoliation and/or the restoration of an intact monolayer caused by migration of neighboring cells. Prior data demonstrates the close correlation of damage area and dead cell exfoliation as two independent measurements

of gastric repair (Aihara et

al., 2018). Due to the lack of

intrinsic fluorescent probes,

only exfoliation was

measured to assess repair in

subsequent experiments

using WT organoids. By

visualizing the nuclei with

Hoechst 33342 (Figure

2.3A), PD to the single cell Figure 2.3 Assessment of dead cell exfoliation to indicate repair during PD in gastric organoid model. A) Representative series of confocal images of WT gastric organoid stained with Hoechst33342 nuclei (blue) before and up to 10 min results in the exfoliation of following single cell PD. Single cell PD occurs in asterisk (shown in yellow). B) Measurement of repair based on exfoliation in control (blue) and EDTA the dead cell towards the supplemented (orange) WT gastric organoids following PD at t=0 min (n = 3). C) Calculated exfoliation (µm) at 10 min based on measurements shown in 6B, as described in Methods (n=3), *p<0.05. lumen.

Under normal conditions, WT reach maximal exfoliation occurs within 10 min while

EDTA supplemented WT gastric organoids show a dampened exfoliation (Figure 2.3B).

At 10 min, WT showed an estimated exfoliation of 8.39 ± 0.90 µm while the presence of

EDTA cause a significant decrease at 0.72 ± 0.32 µm (Figure 2.3C).

To assess repair in gastric organoids derived from fluorescent transgenic mice,

we utilized gastric organoids from YC Nano mice which express a yellow fluorescent

protein (YFP) in the cytoplasm (Oshima et al., 2014). In YC Nano, a single cell within

the gastric organoid was photodamaged and the resulting cell migration and size of

damage area monitored over time. Localized photodamage to part of a single cell

caused prompt loss of cytosolic YFP fluorescence as seen in Figure 2.4A. Over time,

this damage area diminished as cells migrated into the damaged region while the dying

cell exfoliated into the lumen as shown by the reflectance in red (Figure 2.4A). As seen

in Figure 2.4B, within ~10 min, the damage area repairs fully at a repair rate of 0.42 ±

0.07 min-1 (n=7).

Figure 2.4 Assessment of damage area to indicate repair during PD in gastric organoid model. A) Above: Representative series of confocal images of YC Nano gastric organoid structure (YFP, green) before and up to 10 min following single cell PD. Single cell PD occurs in asterisk (shown in yellow). Below: Above confocal images with the addition of confocal reflectance (red) and bright-field. Single cell PD occurs in asterisk (shown in yellow). Following PD, the damaged cell exfoliates into the lumen coinciding with the closure of damaged area by neighboring cells filling in the gap left by departing cell. B) Measurement of repair based on damage area (black) in YC Nano gastric organoids following PD at t=0 min (n = 5).

Results demonstrate that restitution of the gastric organoid is completed promptly following single cell damage and that multiple measurements can report the progression of this event.

2.4.3 Assessing repair in TFF2 deficient gastric organoids

Prior work in vivo has shown that deficiency in trefoil factor 2 (TFF2), a secreted product from mucus cells in the GI tract, results in delayed repair in the stomach (Xue et al., 2010; Xue et al., 2011), other works support that TFF2 is essential in promoting proper repair throughout the GI tract (Babyatsky et al., 1996; McKenzie et al.,

2000; Taupin & Podolsky,

2003; Aihara et al., 2017).

To test if exfoliation and

Lucifer yellow intensity can report repair within TFF2

KO gastric organoids,

Hoechst33342 (nuclei) and

Lucifer yellow (extracellular space) were added to the media of WT and TFF2 KO Figure 2.5 Dead cell exfoliation and Lucifer yellow intensity in TFF2 deficient gastric organoids. A) Comparison of exfoliation in WT (white, n=4) and TFF2 KO (blue, n=4), *p<0.05. B) Measurement of Lucifer yellow intensity (ratio), results gastric organoids and compare before and after (10 min) PD in WT (white, n=4) and TFF2 KO (blue, n=4) gastric organoids. C) Comparison of Lucifer yellow intensities (ratio) shown in 6B at repair was monitored 5 µm from the damage area (both n=4).

following PD. At 10 min following PD, TFF2 KO exhibited a significantly delayed exfoliation of 2.41 ± 0.18 µm (n=4) compared to 9.07 ± 1.20 µm observed in WT (n=4,

*p<0.05) (Figure 2.5A). This data supports in vivo findings that TFF2 deficiency results in delayed repair. To assess permeability and potential leakiness in TFF2 KO gastric organoids, we utilized the approach shown in Figure 2.4D, measuring a line scan in the organoid lumen starting at 5 µm away from the damage site before PD and 10 min following PD (Figure 2.5B). At 5 µm away from the damage site, prior to PD in TFF2

KO (n=4) there was no significant increase in Lucifer yellow intensity within the organoid lumen compared to WT gastric organoids (n=4) (Figure 2.5C). Additionally, no significant difference in luminal Lucifer yellow intensity was observed between before

PD and 10 min after PD in either WT or TFF2 KO gastric organoids (Figure 2.5C). This data suggests that while Lucifer yellow serves as a useful marker of barrier integrity, it may not be the best indicator of repair under certain conditions.

2.4.4 Photodamage induces caspase-activated apoptosis

While the photodamage (PD) model via two-photon microscopy has been established in vivo (Starodub et al., 2008; Xue et al., 2010; Xue et al., 2011; Aihara et al., 2013) and in vitro (Aihara et al., 2018), the type of cell death induced was previously uncharacterized. To address whether PD induces programmed cell death, a fluorogenic caspase 3/7 reagent was added to the medium and PD was induced to single cells and then monitored over time. Caspase 3/7 is a caspase substrate that fluoresces when either activated caspase3 or caspase 7 cleaves the molecule during apoptosis (Figure

2.6).

Caspase 3/7 detection reagent couples a DNA intercalating dye to the activated caspase 3/7 recognition motif DEVD. Following caspase 3/7 activation, caspase 3/7 cleave the DEVD recognition motif causing the DNA stain to intercalate to the nuclei and fluoresce (Yu et al., 2001; Huang et al., 2011).

Figure 2.6 Diagram of Caspase 3/7 green reagent in living cell (left) and apoptotic cell (right). During caspase- activated apoptosis, caspase 3/7 green reagent is cleaved from its DEVD peptide (which is recognized by caspases) and fluoresces upon binding DNA, indicating apoptosis.

We first tested if apoptosis was part of the physiological cell shedding in the absence of PD. As seen in Figure 2.7A, caspase 3/7 fluorescence is only visible in cells that are shed within the gastric organoid, where no PD occurred. The observed caspase

3/7 fluorescence is localized in the same position as reflectance (red) and brightfield showing cells detached from the organoid membrane layer (Figure 2.7A), indicating that these cells have been detached from the membrane. Results suggest that apoptosis occurs under normal conditions of cell shedding and replacement.

To assess whether PD induces caspase-activated apoptosis, caspase 3/7 reagent was added 30 min prior to experimentation and was not visible in viable cells under normal conditions due to the lack of caspase 3/7 activation (Figure 2.7B).

Following PD (Figure 2.7B, C), caspase 3/7 reagent increased in fluorescent intensity, indicating that PD (like normal cell shedding) induces caspase-activated apoptosis

(n=9). To confirm the reliability of caspase 3/7 reagent as an indicator of caspase activated apoptosis, varying concentrations of the pan-caspase inhibitor, Z-VAD-FMK, were added to the media. Following PD, caspase 3/7 fluorescence increased in controls while Z-VAD-FMK dose-dependently prevented induction of fluorescence after PD

(Figure 2.7D, E). Interestingly, while 50, 100 and 150 µM Z-VAD-FMK concentrations prevented induction of caspase 3/7 reagent fluorescence, only the presence of 100 µM and 150 µM Z-VAD-FMK significantly delayed exfoliation (Figure 2.7F, G). Results demonstrate the reliability of caspase 3/7 reagent to identify cell death due to caspase- activated apoptosis and suggest that restitution is dependent on caspase-activated apoptosis.

Figure 2.7 Caspase 3/7 reagent indicates caspase-activated apoptosis in gastric organoids. A) Representative confocal image of unperturbed WT gastric organoid. Arrows indicate cells shed from the organoid membrane; reflectance (red) and caspase 3/7 (green) suggest this cell shedding resulted in cell death, possibly through caspase-activated apoptosis. B) Representative confocal images of Hoechst33342 stained nuclei (blue), caspase 3/7 reagent (green), and confocal reflectance (red) before and following photodamage (PD). Single cell PD occurs in asterisk (shown in yellow). Following PD, caspase 3/7 reagent selectively increases fluorescence in damaged cell. C) Measurement of caspase 3/7 reagent fluorescence intensity over time course in WT gastric organoids following PD at t=0 min (n = 9). D) Representative images, at 1 min following PD, of WT (control) and 50 µM Z-VAD-FMK supplemented gastric organoids. Z-VAD-FMK was added 1 hr prior to experiment. PD indicated by asterisk (shown in yellow). E) Measurement of caspase 3/7 reagent fluorescence intensity over time course in WT (control, n=5), 50 µM Z-VAD-FMK (n=8), 100 µM Z-VAD-FMK (n=8), and 150 µM Z-VAD-FMK (n=5) supplemented gastric organoids following PD at t=0 min. F) Measurement of exfoliation (µm) over time course in WT (control, n=5), 50 µM Z-VAD-FMK (n=8), 100 µM Z-VAD-FMK (n=8), and 150 µM Z-VAD-FMK (n=5) supplemented gastric organoids following PD at t=0 min. G) Comparison of exfoliation distance at 10 min between WT (control, n=5), 50 µM Z-VAD-FMK (n=8), 100 µM Z-VAD-FMK (n=8), and 150 µM Z- VAD-FMK (n=5) supplemented gastric organoids. *p<0.05.

2.5 Discussion

With the advent of the gastrointestinal organoid system by Sato et al. (Sato et al.,

2011), organoids, especially gastric, have been used as an investigative tool to elucidate biological responses in the setting of disease and development, as well as to delineate molecular signaling cascades(Mahe et al., 2013; Schumacher et al., 2015a;

Clevers, 2016; Engevik et al., 2016; Engevik et al., 2018b). While studies involving organoids have become increasingly prevalent, much remains unknown in terms of organoid application. One such area includes assessing repair mechanism within gastric organoids as further insight into in vivo conditions. While in vivo damage models, including ulceration and microlesions (Koo, 1994; Takeuchi et al., 1999; Engevik et al.,

2016; Engevik et al., 2018b), have been well established to study the repair process at a systemic level, in vitro damage models have been limited to cancer cell lines and the use of inhibitors or scratch wound assays to provide more molecular information regarding the signaling cascades involved in gastric repair (Ranta-Knuuttila et al., 2002;

Redlak et al., 2007; Tétreault et al., 2008; Agle et al., 2010). While such studies have provided the bulk of our current knowledge in gastric repair, much remains unknown especially regarding normal native tissue at the epithelial level. This study aimed to determine the cellular response to single cell damage and determine methodology to evaluate repair in a reliable manner across gastric organoid models.

Two-photon induced microlesions have been accepted as a damage model in vivo (Demitrack et al., 2010; Oshima et al., 2014; Clevers, 2016; Engevik et al., 2016;

Engevik et al., 2018b), however the type of cell death induced by this method was previously largely unknown. In this study we show that two-photon damage induces

caspase-activated apoptosis within the gastric organoid system, which is a similar response as seen during natural cell shedding. Caspase-activated apoptosis is an extremely controlled programmed cell death, involving at least five caspases that act either as an initiator or executioner during the apoptosis process (Danial & Korsmeyer,

2004; McIlwain et al., 2013). Caspases -3, -6, and -7 act as executioner caspases, which must be activated via the cleaving of inactive procaspase dimers by the initiator caspase -8 or -9. Interestingly, the pan-caspase inhibitor Z-VAD-FMK was sufficient at all tested concentrations to effectively block caspase-activated apoptosis following PD, however only higher Z-VAD-FMK concentrations effectively blocked restitution. These results suggest that Z-VAD-FMK at lower concentration is less effective in inhibiting the caspase important for cell shedding. While more studies are needed to further investigate the role of caspase-activated apoptosis during gastric repair, the caspase assay gives insight to the controlled mechanism by which repair occurs at the single cell level and supports the use of photodamage as a model that simulates and stimulates natural cell processes that promote cell turnover.

In native tissue, it is essential that gastric epithelial cells maintain close contact to neighboring cells and the substratum to prevent damage from the noxious environment of the stomach. Similar to what is seen in vivo (Xue et al., 2010; Aihara et al., 2013), localized photodamage results in rapid dead cell exfoliation, coincided with the migration of adjacent viable cells to cover the denuded area, sustaining epithelial continuity. The lack of Lucifer yellow entering the lumen following photodamage and during the repair process suggests that the integrity of the epithelium in maintained during restitution. This has also been observed in other gastric organoid models using

fluorescent dextran (Schultheis et al., 1998). Furthermore, unperturbed organoids maintain the integrity of the epithelial barrier and show no sign of basal cell migration.

Lucifer yellow served as a qualitative marker of organoid epithelial integrity in control especially when compared to EDTA supplemented gastric organoids which showed a compromised barrier following PD due to the ability of EDTA to disrupt tight junctions through chelation of calcium (Aihara et al., 2018). EDTA also proved effective in delaying dead cell exfoliation, potentially suggesting the role of extracellular calcium in repair as well as the use of Lucifer yellow as a reliable marker of repair. However, under less severe conditions, such as TFF2 deficiency (ie KO), while delayed repair was observed there was no significant increase in Lucifer yellow intensity within the organoid lumen compared to WT organoids. This suggest TFF2 deficiency does not have a detrimental effect upon tight junctions and cell-cell contact, however TFF2 KO does affect the ability to properly expel damaged or dead cells from the epithelium. Based on this evidence, while Lucifer yellow may serve as an extracellular marker it does not necessarily yield quantitative data to shed more light upon gastric organoid repair.

Nonetheless, the epithelial integrity and proper barrier function observed under normal conditions indicates the organoid system is a reliable in vitro model that can recapitulate certain aspects seen in native tissue.

Gastric repair is a complex process involving both restitution and regeneration.

Gastric restitution is the initiating event for epithelial repair and involves rapid re- establishment of epithelial integrity following injury. This event occurs before regeneration (cell proliferation) or inflammatory responses. Following injury, dead or damaged cells are expelled into the gastric lumen and adjacent viable cells release

bioactive peptides, or motogens, which signal cells to cover the denuded mucosa without proliferation (Svanes et al., 1982; Lacy & Ito, 1984). This process rapidly restores epithelial continuity and barrier function. Gastric restitution is speculated to depend entirely on the epithelial cells themselves as restitution can be demonstrated in vitro in cell lines (Rutten & Ito, 1983; Svanes et al., 1983) and recently in gastric organoids (Aihara et al., 2018). Since the progress of cell exfoliation is a product of the movement of viable cells into the area of damage, we have developed methods to quantify the cell exfoliation as a measure of epithelial repair.

Chapter 3

Comparison of genetically encoded calcium sensors to assess calcium mobilization in gastric organoid repair studies

Chapter 3 Comparison of genetically encoded calcium sensors to assess calcium mobilization in gastric organoid repair studies 3.1 Abstract

Calcium (Ca2+) is a known accelerator for gastric wound repair, however the ability to measure in vivo and in vitro has been limited to Ca2+ sensitive dyes or use of genetically encoded calcium indicators (GECI). While Ca2+ sensitive dyes have proved to be useful, these dyes do not properly load in gastric tissue, making GECIs essential to assess Ca2+ during repair. We previously demonstrated in vivo, using GECI mice transgenic for a fluorescent calcium reporter, that Ca2+ mobilization is essential for gastric epithelial repair. We recently obtained the newly generated GECI transgenic mice expressing Yellow Cameleon (YC) Nano15, which have a brighter fluorescence and a higher affinity as a Ca2+ reporter in comparison with our previously used GECI transgenic mice, YC 3.0. YC sensors utilize Förster Resonance Energy Transfer (FRET) to assess changes in intracellular Ca2+. To compare the two YC systems, we generated gastric organoids from YC Nano and YC 3.0 mice stomach corpus. The criteria used to assess these sensors included organoid size, overall fluorescence intensity, response to damage, noise of fluorescence measures, visibility of cell structure, overall image quality, and changes in FRET/YFP ratio following damage. We also compared use of two-photon excitation (840 nm) versus visible light excitation (458 nm) for optimizing the

FRET signal. In this chapter, we seek to compare these two GECIs to determine (1) which YC sensor provides the most reliable and sensitive Ca2+ reporter and (2) if the in vitro gastric organoid model mimics the Ca2+ mobilization observed in vivo. Current outcomes suggest the usefulness of YC Nano as a more sensitive and reliable sensor

to further investigate intracellular Ca2+ dynamics and elucidate the signaling cascade behind Ca2+-mediated repair.

3.2 Introduction

The use of fluorescence probes to study biological structure and physiology has been long recognized as a powerful tool. Fluorescent probes are molecules chemically attached to aid in the selective detection of another biomolecule such as a protein. The introduction of green fluorescent protein (GFP) in 1962 revolutionized fluorescent microscopy, advancing live cell imaging initially with the ability to selectively tag specific regions of living cells and observe trafficking of specific protein of interest (Shimomura et al., 1962; Prendergast & Mann, 1978; Tsien, 1989; Heim & Tsien, 1996; Renz, 2013).

Protein labeling with fluorescent protein has proved important due to the ability to express fusion proteins in cellular environments with high sensitivity and often non- destructive to cellular function (Tsien, 1989; Sahoo, 2012). Since the introduction of

GFP, several spectral variants, including yellow fluorescent protein (YFP) and cyan fluorescent protein (CFP), have been generated with improved efficiency and enhanced fluorescence brightness (Sahoo, 2012; Renz, 2013). While these fluorescent proteins have been advantageous in self-fluorescent properties and biocompatibility both in vivo and in vitro, often being larger in size than the protein of interest making it difficult to tag certain proteins without affecting either the conformation or function of the targeted protein (Tsien, 1989; Sahoo, 2012).

As research advanced, the fluorescence proteins then were engineered to become biosensors for multicolor live imaging of various processes including cell

division, organelle structure, or even signaling events. Fluorescent proteins has been an advantageous technique for in vivo and in vitro live imaging studies including calcium

(Ca2+) imaging (Swanson et al., 2011; Horikawa, 2015; Perry et al., 2015), pH sensing

(Swanson et al., 2011; Dennis et al., 2012; Georgiev et al., 2019), measurement of chloride (Illsley & Verkman, 1987; Zhou et al., 2012; Arosio & Ratto, 2014), and redox potential (Hanson et al., 2004; Meyer & Dick, 2010; Swanson et al., 2011). The advancements made in fluorescent proteins have allowed for more in-depth studies on intracellular processes.

Calcium is a known accelerator of gastric wound repair (Miller & Henagan, 1979;

Critchlow et al., 1985), however the mechanism by which calcium mobilization affects restitution has yet to be elucidated. While intracellular loading of conventional acetoxy methylester Ca2+ sensitive fluorescent probes, such as fura-2 or fluo-4, have been utilized in several in vitro studies, for reasons unknown these probes fail to load effectively in vivo in normal gastric cells (Aihara et al., 2013). The mechanistic basis of

Ca2+ mobilization in healthy tissue has largely been unexplored due to past limitations of

Ca2+ sensors and difficulty in monitoring intracellular Ca2+ or wound repair in real time.

Recent advancements in high resolution microscopy and genetically encoded Ca2+ indicators (GECI) in mice present the opportunity to monitor Ca2+ mobilization during epithelial repair, heightened by use of gastric organoids which allow the ability to bypass systemic effects of inhibitors or agonists while also reflecting expression similar to native tissue.

Our lab has previously used yellow cameleon (YC) 3.0 transgenic mice to show that endogenous Ca2+ mobilization is required for proper wound repair in vivo (Aihara et

al., 2013). Recently, our lab acquired a newly generated YC transgenic mouse line known as YC Nano15 (YC Nano), which ubiquitously expresses a Ca2+ sensing protein that has been shown in pancreatic studies to be more sensitive to changes in Ca2+ signaling (Oshima et al., 2014). Furthermore, YC Nano has been reported to express brighter fluorescence and a higher affinity Ca2+ reporter compared to earlier YC sensors, including YC 3.0 (Horikawa et al., 2010; Oshima et al., 2014; Horikawa, 2015).

YC Ca2+ sensors allow the use of Förster Resonance Energy Transfer (FRET) to assess changes in intracellular Ca2+ in vivo and in vitro. In FRET-based indicators, fluorescent protein pairs containing a donor and acceptor are linked to visualize molecular interactions within living cells (Arai & Nagai, 2013). YC sensors are a single peptide chain encoding in a linear sequence cyan fluorescent protein (CFP), a portion of the

Ca2+ binding protein calmodulin, a yellow fluorescent protein (YFP) and the M13 portion of myosin light chain (MLC) (Figure 3.1A). In YC sensors, CFP is the donor

(430 nm excitation, 480 nm emission) and YFP is the acceptor protein (517 nm excitation, 528 emission) (Figure 3.1A, B). Overlap between the CFP emission spectrum and the YFP excitation spectra forms the basis for the efficiency of the donor-acceptor pairing. In the presence of Ca2+, the Ca2+-bound calmodulin domain binds to the calmodulin binding peptide, M13, domain of the MLC (Miyawaki et al.,

1997; Horikawa, 2015), which shortens the distance between the proteins and thereby increases Förster resonance energy transfer (FRET) efficiency between CFP and YFP. Importantly FRET does not occur unless the donor and acceptor are within

1-10 nm of each other (Pérez Koldenkova & Nagai, 2013), keeping FRET to a minimum under conditions of low Ca2+. The ratio imaging of YC reduces any artifacts

introduced by motion or change in focus during the live cell imaging process, which may occur with non-ratiometric indicators. In YC Nano, the construct has a modified calmodulin and MLC which theoretically separate the YFP and CFP fluorescent probe by 15 nm from one another in the absence of Ca2+, which theoretically yields a more sensitive and brighter calcium sensor as earlier YC constructs are separated by more than 15 nm (Horikawa et al., 2010).

In the presence of free intracellular Ca2+ in the cell cytosol, the

YC FRET probe undergoes a conformational change that brings the fluorescent proteins closer, allowing the emission of CFP to directly excite the YFP. This YFP fluorescence generated by FRET causes an increase in the FRET channel, such that the FRET/CFP ratiometric measurement can report a change in intracellular Ca2+. In the absence or unchanged intracellular Ca2+ in the Figure 3.1 FRET based genetically encoded calcium indicators. A) Schematic of Förster resonance energy transfer cytosol, no conformational change (FRET) used to measure intracellular calcium. YC mice have genetically encoded calcium indicators where a linked cyan occurs within the FRET probe which fluorescent protein (CFP) and yellow fluorescent protein (YFP) bind calcium and undergo a conformational change that brings CFP and YFP close enough to allow CFP excitation to induce prevents YFP from being excited and YFP emission via FRET. B) Diagram of absorption and emission spectrum of CFP and YFP wavelengths. results in lower YFP levels overall

thereby resulting in an overall FRET/CFP ratio.

By utilizing FRET, we sought to compare organoids of our previously used YC

3.0 transgenic mice and newly acquired YC Nano, which to date has the highest reported binding affinity and reliable reporter of FRET among YC sensors (Horikawa et al., 2010; Oshima et al., 2014; Horikawa, 2015), in order to establish the best approach to assessing calcium mobilization and to dissect further into the mechanism behind calcium mobilization.

3.3 Materials and Methods

3.3.1 Animal husbandry

Experiments used transgenic mice (C57BL/6 background) expressing the Yellow

Cameleon 3.0 (YC 3.0) (Aihara et al., 2013) or Yellow Cameleon Nano15 (YC Nano) calcium sensor fluorescent proteins (Oshima et al., 2014). Pups were genotyped by genomic PCR as described (Schultheis et al., 1998; Bell et al., 1999; Farrell et al., 2002) and used for experimentation at 2-3 months of age. Animals were given standard rodent chow diet and water, both ad libitum. All animal procedures were approved by the

Institutional Animal Care and Use Committee of the University of Cincinnati.

3.3.2 Mouse-derived corpus organoid culture

Gastric organoids were generated from mouse gastric corpus as described (Mahe et al., 2013; Schumacher et al., 2015a; Engevik et al., 2018b). Isolated gastric glands from the corpus were cultured in Matrigel (Corning) diluted 1:1 in Dulbecco's phosphate buffered saline (DPBS, Corning) without Ca2+ and Mg2+ in 8-well Lab-Tek chamber with

coverglass (Thermo Scientific) to grow gastric organoids. Gastric organoids were cultured in a 5% CO2 incubator at 37°C for 3-4 days prior to experiments.

3.3.3 Induction of two-photon laser-induced photodamage

NIR 40x objective lens. To directly excite CFP fluorescence, either Argon laser 458 nm excitation or titanium-sapphire laser (Ti-Sa) 840 nm excitation was used. Resultant fluorescence emission was simultaneously measured in two channels: YFP-FRET

(535-590 nm) and CFP (500-530 nm), along with simultaneous transmitted light and confocal reflectance images (the latter reflecting 840 nm light to show cell structure).

Wavelength selections for Ca2+ imaging were guided by previous work with YC sensors

(Horikawa et al., 2010; Oshima et al., 2014). In all photodamage experiments, after collecting a set of control images, a small rectangle region (≈5 µm2) of a single cell was repetitively scanned at high Ti-Sa laser power (840 nm: 630 mW average) for 500 iterations (equivalent of 10 min). Experiments examined gastric organoids embedded in

Matrigel, located approximately 50-300 μm from the cover glass.

3.3.4 Image analysis

Damaged area was quantified from the time course of images as described (Xue et al., 2010; Xue et al., 2011; Aihara et al., 2013; Aihara et al., 2014) using Image J and/or

Metamorph software (ver. 6.3, Molecular Devices, Downington, PA, USA). The

damaged area was measured as the region of cellular loss of YFP fluorescence in YC

Nano and YC 3.0 gastric organoids. In each experiment, we determined the time point displaying maximal damage area and estimated rates of epithelial restitution starting from this time using a single exponential curve fit to the changing size of damaged area over time (Xue et al., 2010). Best fit values of the rate constant were used as estimates of the rate of repair (units of min−1).

Changes in intracellular Ca2+ were measured under 458 nm and 840 nm excitations in both YC 3.0 and YC Nano gastric organoids. Regions of interest (ROIs) were determined by bright field (see Figure 3.3A) and YFP images to distinguish cellular structure for measurement. Images were collected under either 458 nm or 840 nm excitation, background (laser off) images were subtracted, then pixel intensities of the

YFP image were divided by corresponding CFP image to obtain the FRET/CFP ratio image. FRET/CFP ratio was normalized to a value of 1 in the averaged pre-damage base lines. FRET/CFP ratio intensities were obtained from the ROIs of intact cells adjacent to the damaged area (adjacent), as well as in intact cells 2- 4 cells away from damage site (undamaged or intact).

3.3.5 Statistical analysis

All values were reported from representative experiments as the mean ± standard error of the mean (SEM) from multiple experiments. Statistical significance was determined using unpaired Student’s T-test, or one-way ANOVA with Dunnett’s multiple comparison post-hoc test. A p value of <0.05 was considered significant.

3.4 Results

3.4.1 Organoid size of YC 3.0 and YC Nano is consistent with other organoid models

Organoids from both YC 3.0 and YC Nano showed normal growth patterns and morphology in comparison with previous organoid models observed (Mahe et al., 2013;

Aihara et al., 2014; Oshima et al., 2014). After 6 days of culture, the average diameter for YC Nano was 536.1 ± 49.5

µm (n= 35) while the average diameter of YC 3.0 was 528.9

± 30 µm (n= 35) (Figure 3.2A,

B). There was no significant difference in terms of organoid Figure 3.2 Comparison of YC 3.0 and YC Nano organoid size size, even when compared to at day 5 of culture. A) Brightfield images of YC 3.0 and YC Nano non-fluorescent C57BL/6 WT gastric organoids taken at day 5 (scale bar = 100μm). B) Comparison of organoid size organoids. Results show that diameters of C57BL/6 WT (white, n=34), YC 3.0 (green, n=35) and expression of YC Nano or YC YC Nano (black, n=35) gastric organoids (n=3 mice per sensor). 3.0 fluorescent proteins do not impact gastric organoid growth patterns or morphology, suggesting the presence of these reporters do not affect the gross viability of cells in the organoid.

3.4.2 Quality of imaging and data from YC 3.0 vs YC Nano

Fluorescence excitation under wavelengths of 458 nm or 840 nm excitation was chosen for the most selective laser excitation of CFP by a single or two-photon method based upon previous work (Horikawa et al., 2010; Aihara et al., 2013; Oshima et al.,

2014). Under either excitation, YFP and CFP were detected within both YC sensors.

However, at identical settings of detector sensitivity in the microscope, YC Nano proved brighter compared to the YC 3.0, as seen in Figure 3.3A. Individually, under 458 nm excitation, both YC sensors yielded a brighter and less noisy/grainy image.

Measurement of FRET/CFP ratio in

intact cells under 458nm and 840nm

excitation in YC Nano or YC 3.0

showed relatively constant levels over

time, suggesting stable intracellular

Ca2+ levels (Figure 3.3B).

Normalized average intensities of

CFP and YFP levels were measured in

intact undamaged cells independent of

FRET/CFP ratio measurements to

assess levels in intact organoids and

assess whether the values were stable,

as seen in the FRET/CFP ratio of

undamaged cells in Figure 3.5.

Figure 3.3 Visibility and excitation detection in YC 3.0 and YC Under 458 nm excitation, the YFP Nano. A) Confocal images of intact YC Nano (top) and YC 3.0 (bottom) organoid under brightfield or 458 nm or 840 nm excitation (scale bar = 10μm). B) Timecourse measurement of FRET/CFP and CFP average intensities in YC ratios under 458 nm (left) or 840 nm (right) excitation of YC Nano (top) and YC 3.0 (bottom) intact organoids. Nano ranged within 0.93-1.13 (YFP) and 0.94-1.14 (CFP) respectively (Figure 3.4A). Under 840 nm excitation, the YFP and

CFP average intensities in YC Nano ranged within 0.93-1.12 (YFP) and 0.89-1.12

(CFP) respectively (Figure 3.4B). Under 458 nm excitation, the YFP and CFP average intensities in YC 3.0 organoids ranged within 0.87-1.12 (YFP) and 0.89-1.13 (CFP) respectively (Figure 3.4C). Under 840 nm excitation, the YFP and CFP average intensities in YC 3.0 organoids ranged within 0.90-1.11 (YFP) and 0.84-1.15 (CFP) respectively (Figure 3.4D). Compared to the YC Nano organoid under 840 nm excitation, the CFP values within the YC 3.0 were much more variable. As YFP excitation overlaps with 458 nm, resulting in YFP indirectly being excited by the 458 nm excitation, we utilized Ti-Sa excitation at 840 nm to obtain a more accurate representation of

FRET/CFP ratio.

Under either excitation Figure 3.4 Average intensities of YFP and CFP in YC 3.0 and YC Nano gastric organoids. A) Timecourse measurement of YFP (red) and CFP the YC Nano proved to be the (green) in intact YC Nano gastric organoids under 458 nm excitation. B) Timecourse measurement of YFP (red) and CFP (green) in intact YC Nano gastric organoids under 840 nm excitation. C) Timecourse measurement of most visible in fluorescence at YFP (red) and CFP (green) in intact YC 3.0 gastric organoids under 458 nm excitation. D) Timecourse measurement of YFP (red) and CFP (green) in settings low enough to prevent intact YC 3.0 gastric organoids under 840 nm excitation. photobleaching. Under the same settings, YC 3.0 imaging was dim and it was difficult to fully image the photodamage process as well as distinguish cellular spaces based on fluorescence. These results suggest YC Nano as a more reliable sensor.

3.4.3 Comparison of response to damage in YC 3.0 and YC Nano

Our lab has previously used two-photon photodamage (PD) in vivo and in vitro as a method to target gastric cells (Xue et al., 2010; Xue et al., 2011; Aihara et al., 2013;

Aihara et al., 2018) and optically monitor repair by measuring the restoration of an intact epithelium caused by migration of neighboring viable cells. In both YC sensor organoids, localized photodamage to part of a single cell caused prompt loss of cytosolic

YFP and CFP fluorescence as seen in Figure 3.5A. Over time, this damage area diminishes as neighboring cells migrated into the damaged region (Figure 3.5B).

Consistent with previous findings, closure of the damaged area is essential for complete repair

(Aihara et al., 2018). As described

(Methods), repair was quantified Figure 3.5 Photodamage and repair in YC 3.0 and YC Nano gastric organoids. A) Confocal images of YC Nano (top) and YC 3.0 (bottom) by measuring the damage area gastric organoid structure (merged YFP and CFP channels) under 458 nm excitation before and after photodamage (PD) (scale bar = 10μm). B) size over time (n=4) (Figure Measurement of repair in YC Nano (black) and YC 3.0 (green) gastric organoids following PD at t=0 (n=4). C) Comparison of maximum damage area between YC Nano (n=4, black) and YC 3.0 (n=4, green) within 1 min 3.5B). Over the time-course of following PD. D) Comparison of damage area between YC Nano (n=4) and YC 3.0 gastric organoids (n=4) over time. E) Comparison of rate of repair, calculated based on the damage area overtime between YC Nano 10 min, no statistical difference (black) and YC 3.0 (green) gastric organoids (n=4).

was found in any of the damage/repair metrics between YC 3.0 and YC Nano gastric organoids sensors (Figure 3.5C, D). In both YC sensor organoids, the damage area repairs fully within 10 min, where YC Nano repairs at a rate of 0.45 ± 0.09 min-1 (n=4) and YC 3.0 repaired at a rate of 0.40 ± 0.12 min-1 (n=4) (Figure 3.5E). These results suggest both YC sensors as a reliable organoid model for assessing repair.

3.4.4 Calcium mobilization response of YC 3.0 and YC Nano

YC Nano and YC 3.0 sensors ubiquitously express a fluorescent protein that is a sensitive Ca2+ reporter (Aihara et al., 2013; Oshima et al., 2014; Horikawa, 2015), allowing use of Förster resonance energy transfer (FRET) to measure intracellular Ca2+ via ratiometric imaging (Methods). The spaces within cells directly neighboring the damage site (adjacent) were used for assessment of Ca2+ mobilization, as well as in cells at least 2-4 cells away (undamaged) from the damage site (Figure 3.6A). Upon photodamage of single cells expressing either YC sensor, cells adjacent to the damage site (adjacent) elicit Ca2+ mobilization as measured by FRET ratio (Figure 3.6A, B).

Under 458 nm excitation, FRET/CFP ratios in the adjacent site peaked at 1.2 ± 0.04 in

YC Nano (n=4) and at 1.56 ± 0.22 in YC 3.0 (n=4) (Figure 3.6C). Intact cells

(undamaged) that were 2-4 cells away from the damage site yielded no statistical differences in FRET/CFP ratio overtime. Under 840 nm excitation, following PD the

FRET/CFP ratios in the adjacent site peaked at 1.45 ± 0.04 in YC Nano (n=4) and at 2.0

± 0.28 in YC 3.0 (n=4) (Figure 3.6C), with the undamaged cells yielded no detectable change in FRET/CFP ratio. Under 458 nm excitation, YC 3.0 demonstrated a stronger change in calcium following photodamage compared to the YC Nano (Figure 3.6C).

Considering the probability of 458

nm excitation directly exciting

YFP, as well as the degree of

noise observed in Figure 3 and

Figure 3.4, it is possible that the

increases noticed may not

properly reflect actual changes in

FRET. Under 840 nm excitation,

both YC sensors yielded an

increased FRET/CFP ratio peak

compared to 458 nm excitation.

This suggests that 458 nm may

be directly exciting YFP,

decreasing the FRET/CFP ratio.

Compared to YC 3.0,

Figure 3.6 Assessing calcium mobilization in YC 3.0 and YC Nano under 840 nm excitation, the YC during epithelial repair. A) FRET/CFP ratio median filter image of YC Nano gastric organoid under 458 nm excitation before and 3 min after Nano was significantly less in its photodamage (PD) (scale bar = 10μm). B) Time course measurement of normalized FRET/CFP ratio of neighbor cells adjacent to the damage site (adjacent, red) and intact viable cells 2-4 cells away from the damage site Ca2+ response. Unlike YC 3.0, (undamaged, blue), and two cells away from the damage site (n=4). C) Comparison of maximum FRET/CFP ratio between YC Nano (black) and YC 3.0 (green). YC Nano FRET/CFP ratios were more stable and consistent across the samples as well as presented less noise in the imaging process. Under 840 nm excitation, YC 3.0 showed a greater change in

FRET/CFP values following damage compared to YC Nano; however, YC Nano yielded more stable FRET/CFP ratio levels in undamaged/intact cells compared to YC 3.0

which had varying values. These results suggest that YC Nano under 840 nm excitation as a more suitable sensor for Ca2+ mobilization.

3.5 Discussion

Calcium (Ca2+) signaling affects most aspects of cellular function and life; however, the ability to monitor and measure the intricacies of cellular Ca2+ handling varies across systems. While Ca2+ indicator dyes have proved to facilitate Ca2+ imaging in cultured cells in vitro and some animal models in vivo, these dyes often fail to localize in certain cell subpopulations. Additionally dyes often have poor retention under hours of observation (Horikawa, 2015). Within the stomach, there arise additional difficulties in using traditional Ca2+ sensitive dyes due to its acidic nature (Aihara et al., 2013). Due to these difficulties, we utilized gastric organoids from transgenic mice that contained a

FRET calcium sensor. In previous studies, YC 3.0 transgenic mice were used to assess changes in Ca2+ mobilization in response to damage (Aihara et al., 2013). Due to the recent introduction of an improved YC sensor, we obtained the YC Nano transgenic mice as a potentially more sensitive GECI. Here we report the testing of two Ca2+ sensors to determine the most appropriate choice to assess Ca2+ mobilization during repair of gastric epithelium.

Two-photon PD was utilized to cause focal damage which represents a unique experimental model to study repair especially at the single cellular level, as it allows the induction of microlesions in seconds, ability to control the site of damage, and ability to control initial size of damage. This model has been utilized in various repair studies spanning intestine, pancreatic islets, embryos and stomach (Rocheleau et al., 2004;

Shen et al., 2008; Clark et al., 2009; Xue et al., 2010; Xue et al., 2011; Aihara et al.,

2013). In addition to a localized site of damage, using this method we are able to perform confocal time-lapse microscopy that allows for the quantification of epithelial repair progression.

We established criteria (Table 3.1) to compare gastric organoids from YC 3.0 and YC Nano transgenic mice, in order to test the strength of each YC sensor.

Additionally we tested 458 nm and 840 nm excitation within both YC gastric organoids based upon previous work (Horikawa et al., 2010; Aihara et al., 2013; Oshima et al.,

2014), to confirm the best excitation to observe FRET/CFP changes that accurately reflected indications of Ca2+ mobilization. As shown in Table 3.1, criteria were established and tested to determine the strength of each sensor. The criteria included comparisons of organoid growth, overall imaging quality, overall intensity, visibility of cell structure, direct excitation of YFP,

Table 3.1 Criteria for most reliable and sensitive overall noise of data, response to genetically encoded calcium indicator. damage, and change in FRET/CFP ratio following damage (Table 3.1).

YC 3.0 is an established sensor used both in in vivo and in vitro studies

(Nyqvist et al., 2005; Aihara et al., 2013;

Horikawa, 2015). Similar to what was observed in our organoid growth, YC 3.0 and YC Nano transgenic mice are healthy and the transgenes have no measurable

detrimental effect on the physiological responses of importance for our future study

(Nyqvist et al., 2005; Aihara et al., 2013; Oshima et al., 2014). Within Table 3.1, we further tested the YC sensors fluorescence under either 458 nm or 840 nm excitation.

Cell structure, determined by YFP images, in YC Nano proved most visible under either excitation while at the same settings YC 3.0 structure was difficult to visualize via fluorescence. Furthermore, YC 3.0 had a lower overall fluorescence image quality and overall intensity compared to YC Nano in either excitation. In terms of noise of data as shown in Figure 3 and Figure 3.4, YC Nano under 840 nm excitation was the lowest while YC 3.0 showed moderate to high noise. Based upon excitation and emission information, it is likely that under 458 nm excitation YFP may be directly excited which could result in inaccurate FRET/CFP ratio measurements. This suggests that 458 nm may not be the optimal setting to assess FRET/CFP ratios within either YC sensor. Both

YC sensors responded to PD, with a similar rate of repair occurring within ~10 min similar to what has been observed in other gastric organoid models (Aihara et al.,

2018). Damage to the single cell resulted immediately in an increased FRET/YFP ratio in the cells directly neighboring the damage site in both YC sensors. In either YC sensor, the greatest change was observed under 840 nm.

As a GECI, the YC Nano sensor has been recently generated and reported in studies investigating pancreatic cells (Horikawa et al., 2010; Oshima et al., 2014). YC

Nano is reported to be one of the highest-sensitive GECI and is available in a transgenic model expressing the protein ubiquitously (Oshima et al., 2014), allowing the study of calcium dynamics in any system of interest. This study is the first to the author’s knowledge in utilizing the YC Nano transgenic sensor to establish organoids

and study Ca2+ mobilization during epithelial repair. Based upon our criteria, we determined the YC Nano as the more optimal sensor, due to its bright expression, consistency and low noise in FRET/CFP ratio measurements with and without damage, as well as its resistance to photobleaching. While YC 3.0 yielded significantly higher

FRET/CFP ratio peaks following damage, due to the high variability in intact cellular

YFP and CFP levels, as well as the difficulty in properly visualizing under fluorescence it was deemed as a less reliable sensor to monitor Ca2+ mobilization during repair.

Furthermore, due to the difficulty in measuring the areas for the YC 3.0 as the image quality is not as clear, there is a possibility that these measurements are not true reflections of actual calcium mobilization. Due to these observations, YC Nano was determined to be a more reliable and sensitive indicator in Ca2+ mobilization. Using the

YC Nano derived gastric organoid we identified important Ca2+ signals originating at the leading edge of the damage site. This work significantly advances our ability to assess

Ca2+ mobilization in gastric cells by using a more optimal sensor, allowing for future work investigating into the role of Ca2+ signaling during gastric restitution.

GECIs allow researchers the ability to examine Ca2+ mobilization in ways that are currently not possible or limited in vivo in mice or humans. Based on existing studies

(Engevik et al., 2016), we believe that gastric organoids are the most physiologically relevant model for examining Ca2+ in response to damage and restitution. We speculate that Ca2+ mobilization behaves in a similar fashion in organoids as it does in native tissue; as a result we believe our data has the potential to translate to human health.

Chapter 4

Trefoil factor 2 activation of CXCR4 requires calcium mobilization to drive epithelial repair in gastric organoids

Chapter 4 Trefoil factor 2 activation of CXCR4 requires calcium mobilization to drive epithelial repair in gastric organoids

The majority of data presented in this chapter is currently under resubmission to be published by Engevik, KA, Hanyu, H, Matthis, AL, Zhang, T, Frey, MR, Oshima, Y,

Aihara, E, and Montrose, MH in J of Physiology 2019. Additional data have also been included.

4. 1 Abstract

The stomach mucosa is continually exposed to environmental and physiological stress factors which can cause local epithelial damage. While much is known about gastric wound repair, the stepwise process that characterizes epithelial restitution remains poorly defined. This work seeks to elucidate effectors that drive gastric epithelial repair using a reductionist culture model. To determine the role of trefoil factor

2 (TFF2) and intracellular calcium (Ca2+) mobilization in gastric restitution, gastric organoids were derived from TFF2 knockout mice and yellow cameleon-Nano15

(fluorescent calcium reporter) transgenic mice, respectively. Inhibitors and recombinant protein were used to determine the upstream and downstream effectors of gastric restitution following photodamage (PD) to single cells within the gastric organoids.

Single cell PD resulted in parallel events of dead cell exfoliation and migration of intact neighboring cells to restore a continuous epithelium in the damage site. Under normal conditions following PD, Ca2+ levels increased within neighbor migrating cells, peaking at ~1 min, suggesting localized Ca2+ mobilization at the site of cell protrusion/migration.

TFF2 KO organoids exhibit delayed repair, however this delay can be rescued by addition of exogenous TFF2. Inhibition of epidermal growth factor receptor (EGFR),

ERK1/2, or a TFF2 receptor, chemokine C-X-C receptor 4 (CXCR4), resulted in significant delay and dampened Ca2+ mobilization. Inhibition of sodium hydrogen exchanger 2 (NHE2) caused significant delay but did not affect Ca2+ mobilization. A similar delay was observed in NHE2 KO organoids. In TFF2 KO gastric organoids, addition of exogenous TFF2 in the presence of EGFR or CXCR4 inhibition was unable to rescue repair. Our work demonstrates that intracellular Ca2+ mobilization occurs within gastric epithelial cells adjacent to the damage site to promote repair by mechanisms that involve TFF2 signaling via CXCR4, and activation of EGFR and

ERK1/2. Furthermore NHE2 is shown to be important for efficient repair, and to operate via a mechanism either downstream or independent of calcium mobilization.

4.2 Introduction

Gastric epithelial barrier integrity and proper repair of a disrupted barrier are essential functions to sustain this primary barrier that protects the inner body from noxious contents of the stomach (Niv & Banic, 2014). When encountering either small or extensive epithelial damage, cell migration promotes rapid re-establishment of epithelial integrity as the initial response of epithelial repair. In the presence of more severe damage, this restitution event occurs before regeneration (cell proliferation) or inflammatory responses. During gastric restitution, dead or damaged cells are expelled into the stomach lumen and adjacent viable cells release bioactive peptides that act as motogens to signal cells to cover the denuded mucosa without proliferation (Svanes et

al., 1982; Lacy & Ito, 1984). The mechanism of restitution is speculated to be intrinsic to the epithelial cells themselves as the process can be demonstrated in vitro in cell lines (Rutten & Ito, 1983; Svanes et al., 1983; Kim et al., 2012; Wang et al., 2012), but the regulation and coordination of this multi-cellular process is poorly understood both in vivo and in vitro. Various factors have been shown in vivo and/or in vitro to influence gastric epithelial restitution including Ca2+, trefoil factor peptides (TFFs), and epidermal growth factor receptor (EGFR) (Hansson et al., 1990; Furukawa et al., 1999; Nie et al.,

2003; Yang et al., 2006; Xue et al., 2010; Xue et al., 2011; Aihara et al., 2013; Aihara et al., 2018).

Ca2+ is a ubiquitous second messenger that influences multiple cellular processes, including mucus secretion and cell migration in various cell types (Belkacemi et al., 2005; Schreiber, 2005; Wei et al., 2008; Aihara et al., 2014; Xie et al., 2017). In vivo, gastric damage elicits increased intracellular and extracellular Ca2+ (Takeuchi et al., 1985; Koo, 1994; Takeuchi et al., 1999) and both are necessary for proper gastric wound repair (Aihara et al., 2013). Inhibition of intracellular Ca2+ release or uptake significantly prevents cell migration following wounding in cultured rabbit gastric cells

(Ranta-Knuuttila et al., 2002). While studies point to the overarching role of endogenous

Ca2+ in gastric epithelial repair, little is known about the upstream signaling to regulate

Ca2+ mobilization.

Another known factor involved in gastric restitution is the motogenic TFF peptide family. TFFs play an important role within the gastrointestinal (GI) mucosal barrier throughout the GI tract (Lefebvre et al., 1993; Nie et al., 2003; Aihara et al., 2017). In epithelial cell culture models, TFFs have been shown to promote cell migratory and anti-

apoptotic activities (Kinoshita et al., 2000; Taupin & Podolsky, 2003; Hoffmann, 2005), thereby identifying potential roles in mediating mucosal repair. In the stomach of both rodents and humans, TFF2 is abundantly secreted from the stomach mucous neck cells

(Hoffmann, 2005; Aihara et al., 2017). TFF2 deficient (TFF2-/-) mice exhibit delayed gastric repair in vivo (Farrell et al., 2002; Xue et al., 2010; Xue et al., 2011; Aihara et al.,

2016); functional assays suggest that C-X-C chemokine receptor 4 (CXCR4) acts as a

TFF receptor in vitro and in vivo (Dubeykovskaya et al., 2009; Xue et al., 2011). During ulcer healing, in vivo epithelial levels of CXCR4 and TFF2 are increased, and addition of exogenous TFF accelerates the healing process (Poulsen et al., 1999; Xu et al., 2013).

TFF2 is also increased in response to Helicobacter pylori infection or severe damage caused by repetitive administration of NSAIDS, and this TFF2 upregulation can precede changes in other growth factors, including EGF (Konturek et al., 1998; Chen et al.,

2018).

Epidermal Growth Factor (EGF) is another peptide produced by the gastric mucosa (Wright et al., 1990), and EGF receptor (EGFR) is present in gastric epithelial cells (Mori et al., 1987; Ménard & Pothier, 1991; Chen et al., 2001). EGF stimulates gastric epithelial cell migration and accelerates wound healing acting via EGFR and

ERK1/2 signaling both in vivo and in vitro models (Tarnawski & Jones, 1998; LI et al.,

2003; Tarnawski & Ahluwalia, 2012). There is a potential link between TFF2/CXCR4 and EGFR. In gastric cancer cell lines, CXCR4-EGFR cross-talk has been shown to promote cell migration (Guo et al., 2007; Cheng et al., 2017). Additionally, it has also been reported that TFF2 can trigger phosphorylation of EGFR in HT29 colon cancer cells (Kinoshita et al., 2000; Rodrigues et al., 2003; Kosriwong et al., 2011). However, it

is not known if such interactions between TFF2/CXCR4 and EGFR occur outside the setting of cancer cell lines.

Evaluating the epithelial signaling cascade associated with gastric restitution in vivo is difficult. Only a limited number of inhibitors and agonists are suitable for in vivo studies, and the tools for manipulating and monitoring intracellular calcium are less precise in vivo. The organoid culture system allows for the growth and differentiation of primary, normal epithelial cells from mouse tissue (Schlaermann et al., 2014; Bartfeld et al., 2015; Aihara et al., 2018). Gastric organoids contain all epithelial cell types of native tissues (Bartfeld et al., 2015); we have previously shown gastric organoids provide a unique reductionist model system for examining the molecular mechanisms of restitution in the gastric epithelium (Aihara et al., 2018). Using gastric organoids from normal and mutant mice, we seek to evaluate involvement of TFF2, CXCR4 and EGFR in calcium-dependent restitution of gastric damage. Our work demonstrates that a novel convergence of the TFF2, EGFR and Ca2+ signaling pathways is essential for gastric epithelial restitution.

4.3 Materials and Methods

4.3.1 Animal husbandry

Experiments used TFF2 knockout (KO) (C57BL/6 background) mice (Xue et al.,

2011), NHE2 knockout (KO) (FVB/N background) mice (Xue et al., 2011), or transgenic mice (C57BL/6 background) expressing the Yellow Cameleon-Nano15 (YC Nano) Ca2+ sensor fluorescent proteins (Oshima et al., 2014). For experiments examining TFF2 KO

(-/-) and NHE2 KO (-/-) genotypes, wild-type controls were composed of +/+ genotypes from the same colony. Pups were genotyped by genomic PCR as previously described

(Schultheis et al., 1998; Bell et al., 1999; Farrell et al., 2002) and used for experimentation at 2-4 months of age. Animals were given standard rodent chow diet and water, both ad libitum. All animal procedures were approved by the Institutional

Animal Care and Use Committee of the University of Cincinnati.

4.3.2 Mouse-derived corpus organoid culture

Gastric organoids were generated from mouse gastric corpus as described

(Mahe et al., 2013; Schumacher et al., 2015a; Engevik et al., 2018b). Isolated gastric epithelium from the corpus was cultured in Matrigel (Corning) diluted 1:1 in Dulbecco’s

Phosphate-Buffered Saline (DPBS, Corning) without Ca2+ and Mg2+ in 8-well or 2-well

Lab-Tek chamber with coverglass (Thermo Scientific) to grow gastric organoids. Gastric organoids were cultured in a 5% CO2 incubator at 37°C for 3-4 days prior to experiments.

4.3.3 Induction of two-photon laser-induced photodamage

NIR 40x objective lens. In some experiments, gastric organoids were pre-incubated for

30 min with the DNA stain Hoechst 33342 (10 μg/ml, Invitrogen) to visualize cellular

nuclei. In experiments intended for analysis of damage area and cell exfoliation in YC

Nano gastric organoids, images of Hoechst 33342 (titanium-sapphire laser [Ti-Sa] excitation 730 nm, emission 435–485 nm) and YFP (excitation 514 nm, emission 535–

590 nm) in the gastric organoid were collected simultaneously with transmitted light and a confocal reflectance image (reflecting 730 nm light to show cell/tissue structure). In

TFF2 and NHE2 WT and KO gastric organoids, images of Hoechst 33342 were collected simultaneously with transmitted light and confocal reflectance images, using wavelengths described above. For assessing intracellular Ca2+ changes in YC Nano gastric organoids, images of YFP-FRET (Ti-Sa excitation 840 nm, emission 535-590 nm) and CFP (Ti-Sa excitation 840 nm, emission 500-530 nm) were collected simultaneously with a transmitted light image. Wavelength selections for Ca2+ imaging were guided by previous work with YC sensors (Horikawa et al., 2010; Oshima et al.,

2014). In all photodamage experiments, after collecting a set of control images, a small rectangle region (≈5 µm2) of a single cell was repetitively scanned at high Ti-Sa laser power (730 or 840 nm: 630 mW average) for 500 iterations (requiring ≈3 s).

Experiments examined gastric organoids embedded in Matrigel, located approximately 100-300 μm from the cover glass. In some cases, BAPTA-AM (50 μM,

Calbiochem) was applied to medium and incubated for at least 30 min prior to experiments. Inhibitors were pre-incubated at least 1 hr prior to experimentation to assure equilibration in Matrigel, and were kept in the medium during experiments.

Inhibitory reagents included: AMD3100 (1 μM, Sigma), AG1478 (200 nM, Cayman

Chemical), FR180204 (10 μM, Tocris), and Hoechst 694 (100 μM, gift from Dr. H.J.

Lang, Sanofi-Aventis, Frankfurt, Germany). Final DMSO concentration in experiments

was <0.1 %. Solvent control groups contained 0.1 % DMSO added to medium. Vehicle

control groups contained either

0.1% DMSO, ddH2O, or dPBS

added to the medium; vehicle was

dependent on the solution the

inhibitors used were constituted in.

Concentrations were determined

based upon prior in vitro studies Figure 4.1. Assessment of potential cytotoxic effect of inhibitors within the gastric organoid model. Comparison of cytotoxicity within intact WT control and inhibitors following 1 hr incubation, using (Chen et al., 2002; Hurst et al., Live/DeadTM Viability/Cytotoxicity kit. Control (n=7); Hoe694 (n=6); AG1478 (n=7); AG1478 (n=5); EDTA (n=7). 4mM EDTA (which is 2008; Aihara et al., 2018) or shown toxic to gastric organoids) was used as a positive control. #p<0.05 vs control. in preliminary experiments to have no observed toxicity as measured by Live/DeadTM Viability/Cytotoxicity kit

(ThermoFisher Scientific) (Figure 4.1). EDTA (4mM, Fluka) was added for 1 hr incubation to confirm the reliability of the Live/DeadTM Viability/Cytotoxicity kit

(ThermoFisher Scientific).

4.3.4 Microinjection

For rescue experiments in TFF2 and NHE2 KO gastric organoids, recombinant human TFF2 (rTFF2; 40µM stock in DPBS; R&D Systems) was microinjected using

methods previously described (Engevik et al., 2018b). Gastric organoids (~400-500µm diameter) were injected with 9 nL rTFF2 40µM stock for an estimated final rTFF2 concentration of 400nM. In rescue experiments utilizing inhibitors, rTFF2 was microinjected following 1 hr pre-incubation with inhibitors. Control vehicle TFF2 KO or

NHE2 KO gastric organoids were microinjected with 9 nL of dPBS.

4.3.5 Image analysis

Damaged area (units of µm2) was quantified from the time course of images as described (Xue et al., 2010; Xue et al., 2011; Aihara et al., 2013; Aihara et al., 2014) using Image J and/or Metamorph software (ver. 6.3, Molecular Devices, Downington,

PA, USA). The damaged area was measured as the region of cellular loss of YFP fluorescence in YC Nano gastric organoids. In each experiment of YC Nano gastric organoids, we determined the time point displaying maximal damage area and estimated rates of epithelial restitution starting from this time with a single exponential curve fit to the size of damage area over time (Xue et al., 2010; Aihara et al., 2018).

Best fit values of the rate constant were used as estimates of the rate of repair (units of min−1). Additionally, movement of nuclei of the damaged cell was traced and exfoliation

(units of µm) was measured as the maximum distance of the dead cell nuclear movement at 20 min following photodamage. This time point was selected as it allowed for observation of delayed exfoliation after addition of inhibitors. Changes in intracellular

Ca2+ were measured as FRET/CFP ratio using YC Nano gastric organoids. Background images were subtracted from FRET-YFP and CFP images, the resultant images were divided on a pixel-by-pixel basis to calculate the FRET/CFP ratio image. All time course

ratio images were then normalized to the averaged pre-damage baseline images.

Regions of interest (ROIs) were determined by transmitted light and 514 nm excited

YFP images to define cellular structures for whole cell and lateral region measurements.

4.3.6 Statistical analysis

All values are reported from experiments as the mean ± standard error of the mean (SEM) from ‘n’ organoid experiments. Statistical significance was determined using unpaired Student’s T-test, or one-way ANOVA with Dunnett’s multiple comparison post-hoc test. A p value of <0.05 was considered significant.

4.4 Results

4.4.1 Organoids as a model of gastric restitution

Our lab has previously used two-photon photodamage in vivo and in vitro as a method to target individual gastric cells (Xue et al., 2010; Xue et al., 2011; Aihara et al.,

2013; Aihara et al., 2018) and optically monitor repair by measuring cell exfoliation and/or the restoration of an intact epithelium caused by migration of neighboring cells.

Recently we have introduced this approach to gastric organoids (Aihara et al., 2018). In

YC Nano gastric organoid, localized photodamage to part of a single cell nucleus

(stained by Hoechst 33342) caused prompt loss of cytosolic YFP fluorescence as seen in Figure 4.2A. Over time, this damage area diminished as neighboring cells migrated into the damaged region (Figure 4.2A). Consistent with our recent findings (Aihara et

al., 2018), both exfoliation of damaged cell(s) and closure of damaged area are essential for complete repair (Aihara et al., 2018). As described (Methods), these parallel events can be quantified by measuring the damage area size and/or the distance of the damaged cell nuclei movement (exfoliation) over time (n=7) (Figure

4.2B). Within ~10 min, the damage area repairs fully (rate of repair 0.42 ± 0.07 min-1, n=7) and maximal nucleus exfoliation is observed (exfoliation distance 9.17 ± 1.45 µm, n=7). Results demonstrate that restitution of the gastric organoid is completed promptly following single cell damage and that multiple measurements can report the progression of this event.

Figure 4.2. Assessment of repair in photodamage organoid model. A) Above: series of confocal images of YC Nano gastric organoid with Hoechst 33342 (red) stained nuclei and gastric organoid structure (YFP, green) before and up to 10 minutes following single cell photodamage (PD). Single cell PD occurs in rectangle area (shown in yellow). Below series of representative illustrations demonstrate measurements taken over time of the damage area and exfoliation of damaged nuclei. Following PD, the damaged cell exfoliates into the lumen coinciding with the closure of damaged area by neighboring cells filling in the gap left by departing cell. B) Measurement of repair based on damage area (black) and exfoliation (red) in YC Nano gastric organoids following PD at t=0 min (n = 7).

4.4.2 Calcium is required for epithelial wound repair in gastric organoids

To assess intracellular Ca2+ mobilization during the epithelial repair process gastric organoids were generated from transgenic YC Nano mice which ubiquitously express a

sensitive fluorescent

Ca2+ reporter (Oshima et al., 2014), allowing use of Förster resonance energy transfer (FRET) to measure intracellular

Ca2+ via ratiometric imaging (Methods).

Based upon transmitted light and YFP images, cellular boundaries were determined (Methods) and used for Figure 4.3. Comparison of intracellular calcium mobilization in cells near the site of damage. Fluorescence of YC Nano gastric organoids was imaged over time, before assessment of Ca2+ and after photodamage (PD). In time courses, PD occurred at t=0 min. A) Confocal FRET (red)/CFP (green) fluorescent merged image of YC Nano gastric organoid at t= 4 min after PD. Representative color outlines refer to area used to measure whole cellular mobilization within intact calcium levels in intact cells adjacent to damage (neighbor, blue), intact cells one cell space away from damage (1 cell away, red), and intact cells two cell spaces away from cells (Figure 4.3A). damage site (2 cells away, purple). Both sides of the damage site were measured and averaged. B) Time course measurement of normalized FRET/CFP ratio from the 3 cellular regions indicated in A (n=4). Cells adjacent to the damage site show greatest Upon photodamage to calcium mobilization after damage. C) Confocal FRET/CFP ratio fluorescent merged image and FRET/CFP ratio image of YC Nano gastric organoid before and 3 min after PD. Representative color outlines refer to area used to measure intracellular calcium single cells in YC- Nano levels in lateral membrane region in intact cell adjacent to the damage site (red) and intact whole cell adjacent to damage site (blue). D) Time course measurement of organoids as shown in normalized FRET/CFP ratio data from regions indicated in C (n=4). The measurement of damage area (gray) is also shown to report time course of repair. Figure 4.3, cells adjacent to the damage site demonstrate Ca2+ mobilization as reported by FRET/CFP ratio (Figure 4.3B, neighbor). Ca2+ mobilization peaked at 0.75 ± 0.30 min and

dissipated by 4.4 ± 1.0 min (n= 7). As shown in Figure 4.3B, maximal Ca2+ mobilization was greater within the cells directly neighboring the damage site versus cells 1 or 2 cell positions away from the damage site (p<0.05). We also tested for subcellular heterogeneity of Ca2+ mobilization within the cells neighboring the damage. In addition to the whole cell measurement as shown in Figure 4.3B, the sub-cellular lateral membrane region directly adjacent to the damage site was measured separately to assess changes in FRET/CFP ratio (Figure 4.3C). In cells neighboring the damage,

Ca2+ within the lateral membrane region mobilized with a similar time course as the whole cell (Figure 4.3D). However, the lateral membrane region showed a significantly greater maximal FRET/CFP ratio change (1.43 ± 0.04, n=4) versus the whole cell measurement (1.18 ± 0.07, n=4, p<0.05). Therefore, the lateral membrane region was measured routinely as a more sensitive indicator of Ca2+ mobilization in all subsequent experiments.

To confirm the importance of intracellular Ca2+ mobilization in gastric restitution,

BAPTA/AM was applied 30 min prior to photodamage to chelate intracellular Ca2+

(Figure 4.4). BAPTA/AM significantly blocked repair (Figure 4.4A, B); the damage area remaining at 10 min in BAPTA-treated organoids (76.27 ± 24.79 µm2, n=4) was significantly larger than in control organoids (2.3 ± 1.7 µm2, n=4, p<0.05) and the corresponding repair rate (Methods) of 0.11 ± 0.04 min-1 for BAPTA-treated organoids was significantly reduced compared to 0.36 ± 0.04 min-1 observed in control gastric organoids (both n=4, p<0.05). Addition of BAPTA/AM significantly blunted Ca2+ signaling within cells adjacent to the damage site in YC- Nano organoids, where the control FRET/CFP ratio peak was 1.49± 0.04 compared to a FRET/CFP ratio peak of

1.14 ± 0.01 in the presence of BAPTA/AM (both n=4, p<0.05) (Figure 4.4C, D). This data indicates that the FRET/CFP ratio measurements reflect intracellular Ca2+ levels, as incubation with BAPTA/AM effectively diminishes the mobilization of free Ca2+ after damage. These results further demonstrate that intracellular

Ca2+ mobilization is necessary for repair within the gastric organoid model.

Figure 4.4. Effect of intracellular calcium chelation on repair and calcium mobilization. Fluorescence of YC Nano gastric organoids was imaged over time. BAPTA/AM (50 µM) was added to organoid medium 30 min prior to experimentation. In time courses, PD occurred at t=0 min. A) Damage area measured in control (black) and BAPTA/AM supplemented gastric organoids (green) over time (n=4). B) Comparison of rate of repair between control (black) and BAPTA/AM supplemented gastric organoids (green) (n=4, *p<0.05). C) Measurement of normalized FRET/CFP ratio of lateral membrane region of cells adjacent to the damage site comparing control (black) and BAPTA/AM supplemented gastric organoids (green). D) Comparison of the maximum FRET/CFP ratio from panel C between control (black) and BAPTA/AM (green) gastric organoids (n=4, *p<0.05).

4.4.3 TFF2 receptor CXCR4 acts upstream of calcium mobilization and is involved in gastric restitution

Epithelial damage is known to elicit the release of TFF2, which acts via CXCR4 within the gastric epithelium (Xue et al., 2010) and in immune cells (Dubeykovskaya et al., 2009). In order to determine if epithelial CXCR4 was involved in gastric organoid

restitution, the CXCR4 inhibitor AMD3100 was added to YC Nano gastric organoids. At

10 min, whereas control gastric organoids exhibited a 5.8 ± 3.9 µm2 damage area and repair rate of 0.41 ± 0.05 min-1, organoids treated with 1 µM AMD3100 displayed a 56 ±

18 µm2 damage area and significantly delayed repair rate of 0.20 ± 0.07 min-1 (n=4, p<0.05) (Figure 4.5A, B). A parallel examination of Ca2+ mobilization revealed that

CXCR4 inhibition significantly

blunted Ca2+ mobilization from

1.43 ± 0.04 FRET/CFP ratio

peak in control to 1.17 ± 0.03

FRET/CFP ratio peak in cells

adjacent to the damage site

(n=4, p<0.05) (Figure 4.5C,

D). Results indicated that

CXCR signaling and CXCR4-

mediated repair involves Ca2+

mobilization.

Figure 4.5. Effect of CXCR4 inhibition on repair and calcium mobilization. Fluorescence of YC Nano gastric organoids was imaged over time. Where indicated, AMD3100 (1 µM) was added to organoid medium 1 hr prior to experimentation. In time courses, PD occurred at t=0 min. A) Damage area measured in control (black) and AMD3100 supplemented gastric organoids (red) (n=4). B) Comparison of rate of repair between control (black) and AMD3100 supplemented gastric organoids (red) (n=4,*p<0.05). C) Measurement of normalized FRET/CFP ratio of lateral membrane region of cells adjacent to the damage site comparing control (black) and AMD3100 supplemented gastric organoids (red). D) Comparison of the maximum FRET/CFP ratio from panel C between control (black) and AMD3100 (red) gastric organoids (n=4, *p<0.05).

4.4.4 TFF2 action requires CXCR4 and calcium mobilization acting downstream during gastric restitution

Prior data show the close correlation of damage area and dead cell exfoliation as two independent measurements of gastric repair (Aihara et al., 2018). Due to the lack of intrinsic fluorescent probes, only exfoliation was measured to assess repair in subsequent experiments using TFF2 WT and KO organoids. In contrast to WT organoids, which exhibited an exfoliation distance of 8.81 ± 0.70 µm (n=7) at 20 min post-injury, exfoliation was significantly diminished in WT organoids treated with

AMD3100 (1.33 ± 0.35 µm, n=6) or BAPTA/AM (1.52 ± 0.24 µm, n=4) (p<0.05) (Figure

4.6A). Compared to WT, TFF2 KO organoids also exhibited a significant reduction of exfoliation at 2.11 ± 0.27 µm (n=10, p<0.05). Treatment of TFF2 KO organoids with

AMD3100 (1.33 ± .35 µm, n=6) or BAPTA/AM (1.31 ± 0.31 µm, n=4) did not alter the already compromised exfoliation. However, the delayed exfoliation observed in TFF2

KO was rescued by microinjection of exogenous rTFF2 into the organoid lumen (8.55 ±

0.94 µm, n=10, p<0.05) (Figure 4.6A). This rescue was not significantly different whether exogenous rTFF2 (400nM) was added to the organoid medium (7.96 ± 0.68

µm, n=4) or microinjected (Figure 4.6B). Due to limited availability of rTFF2, gastric organoids were microinjected for this study. Exogenous rTFF2 was unable to rescue the exfoliation in the presence of AMD3100 (2.22 ± 0.29 µm, n=5) (Figure 4.6A).

CXCR4 inhibition altered Ca2+ mobilization (as shown in Figure 4.3) and separately prevented rTFF2 action during repair, but the link between these two outcomes is unclear. To directly test if TFF2 action is dependent upon Ca2+ mobilization, BAPTA/AM was added to the media in the presence of gastric organoids

microinjected with rTFF2. Incubation with BAPTA/AM prevented the rTFF2 rescue (1.30

± 0.32 µm, n=8, p<0.05) (Figure 4.5). These results indicate that TFF2 action requires

Ca2+ mobilization to promote the repair process.

Figure 4.6. Comparison of exfoliation within WT and TFF2 KO organoids with and without treatments. A) Results from WT and TFF2 KO gastric organoids imaging over time, measuring movement of fluorescent nuclei (Hoechst 33342 stain) after PD. PD occurred at t=0 min. WT and TFF2 KO gastric organoids were treated with AMD3100 (1µM) for 1 hr or BAPTA/AM (50µM) for 30 min before PD as indicated. rTFF2 was microinjected into the lumen of organoids 30 min before study (see Methods). Exfoliation was determined based on maximum distance of damaged nuclei into gastric organoid lumen over 20 min. Vehicle, (WT control, n=7; TFF2 KO control, n=10; TFF2 + rTFF2 Control, n=10), AMD3100 (WT, n=6; TFF2 KO, n=6; TFF2 KO + rTFF2, n=5), BAPTA/AM (WT, n=4; TFF2 KO, n=4; TFF2 KO + rTFF2, n=8). *p<0.05 vs WT vehicle, #p<0.05 vs rTFF2 treatment in TFF2 KO. B) Comparison of exfoliation between rTFF2 microinjection (400nM final concentration, n=7) or rTFF2 addition to the media (400nM final concentration, n=6) in TFF2 KO organoids.

4.4.5 EGFR acts upstream of calcium mobilization and is involved in gastric restitution

In renal, ovarian, and colonic cancer cells, TFF2 and/or CXCR4 have been shown to interact with or activate EGFR (Rodrigues et al., 2003; Guo et al., 2007;

Kosriwong et al., 2011). Since EGFR has been implicated separately in epithelial wound healing (Hansson et al., 1990), YC–Nano organoids were treated with an EGFR inhibitor (AG1478, 200 nM) to test the role of EGFR in gastric restitution (Figure 4.7). At

10 min, compared with control gastric organoids which exhibited a damage area of 1.0 ±

0.5 µm2 and repair rate of 0.43 ± 0.05 min-1, EGFR inhibition significantly delayed epithelial repair with a damage area of 42 ± 26 µm2 (p<0.05) and repair rate of 0.25 ±

0.02 min-1 (p<0.05 (Figure 4.7A, B). Furthermore, EGFR blockade significantly blunted the maximal FRET/CFP ratio peak from 1.32 ± 0.02 in control to 1.04 ± 0.01 (n=4, p<0.05) (Figure 4.7C, D). These results suggest that EGFR promotes Ca2+ mobilization and gastric restitution.

Figure 4.7. Effect of EGFR inhibition upon repair and calcium mobilization. Fluorescence of YC Nano gastric organoids imaged over time. AG1478 (200 nM) was added to organoid medium 1 hr prior to experimentation. In time courses, PD occurred at t=0 min. A) Damage area measured in control (black) and AG1478 supplemented gastric organoids (orange) (n=4). B) Comparison of rate of repair between control (black) and AG1478 supplemented gastric organoids (orange) (*p<0.05). C) Measurement of normalized FRET/CFP ratio of lateral membrane region of cells adjacent to the damage site comparing control (black) and AG1478 supplemented gastric organoids (orange). D) Comparison of the maximum FRET/CFP ratio from panel C between control (black) and AG1478 supplemented gastric organoids (orange) (n=4, *p<0.05).

TFF2 KO organoids were then used to test if the EGFR is a potential downstream effector of TFF2/CXCR4. Addition of AG1478 caused a significant delay in exfoliation (1.76 ± 0.41 µm versus 8.18 ± 0.35 µm in control, n=6, p<0.05) (Figure 4.8).

However, in TFF2 KO organoids, addition of AG1478 (1.56 ± 0.89 µm, n=5) had no additive effect on exfoliation compared to vehicle (1.56 ± 0.31 µm, n=8). Furthermore, addition of rTFF2 significantly rescued exfoliation in TFF2 KO (8.87 ± 0.82 µm, n=8,

p<0.05). However, in the presence of AG1478, rescue by rTFF2 was significantly prevented (2.30 ± 0.55 µm, n=5, p<0.05). Together these results suggest that EGFR acts downstream of TFF2/CXCR4 in the repair pathway. As both receptors are necessary to stimulate Ca2+ mobilization and promote gastric restitution, these data suggest both CXCR4 and EGFR may be acting via the same Ca2+ mobilizing signaling pathway during repair.

Figure 4.8. Comparison of exfoliation within EGFR inhibited WT and TFF2 KO organoids. Results from imaging of WT and TFF2 KO organoids over time; measuring movement of fluorescent nuclei (Hoechst 33342 stain) after PD. Some organoids were treated with AG1478 (200nM) as indicated. rTFF2 was microinjected into the lumen of organoids before study. Exfoliation was determined based on maximum distance of damaged nuclei into gastric organoid lumen over 20 min. Vehicle (WT control, n=6; TFF2 KO control, n=8; TFF2 KO +rTFF2, n=8), AG1478 (WT, n=6; TFF2 KO, n=5; TFF2 KO + rTFF2, n=5). *p<0.05 vs WT vehicle, #p<0.05 vs rTFF2 treatment in TFF2 KO.

4.4.6 ERK1/2 is necessary for the repair process, acting upstream of calcium mobilization

CXCR4 and EGFR both act via ERK1/2 signaling in various systems (LI et al.,

2003; Billadeau et al., 2006; Zimmerman et al., 2011). To test for a role of ERK1/2 within our organoid model, an ERK1/2 inhibitor (10 µM FR180204) was added to YC

Nano gastric organoids. In contrast to control gastric organoids that exhibited a fully repaired space of 0

µm2 damage area at

10 min and repair rate of 0.45 ± 0.04 min-1, organoids treated with

FR180204 displayed a 34.9 ± 2.6 µm2 damage area at 10 min and repair rate of 0.21 ± 0.03 min-1

(Figure 4.9A, B)

(both n=4, p<0.05).

Examination of Ca2+ Figure 4.9. Effect of ERK1/2 inhibition on repair and calcium mobilization. Fluorescence of YC Nano gastric organoids imaged over time. Where indicated, FR180204 (10 µM) was added to organoid medium 1 hr prior to experimentation. In time courses, PD mobilization by occurred at t=0 min. A) Damage area measured in control (black) and FR180204 supplemented gastric organoids (purple) (n=4). B) Comparison of rate of repair between FRET/CFP ratio in control (black) and FR180204 supplemented gastric organoids (purple) (*p<0.05). C) Measurement of normalized FRET/CFP ratio of lateral membrane region of cells adjacent to the damage site comparing control (black) and FR180204 supplemented gastric gastric organoids organoids (purple). D) Comparison of the maximum FRET/CFP ratio from panel C between control (black) and FR180204 supplemented gastric organoids (purple) (n=4, *p<0.05).

revealed that FR180204 dampened Ca2+ mobilization in the cell adjacent to the damage site (1.13 ± 0.02, compared to control 1.33 ± 0.03, both n=4, p<0.05) (Figure 4.9C, D).

These results indicate that ERK1/2 operates upstream of Ca2+ mobilization pathways during repair.

4.4.7 Sodium hydrogen exchanger 2 acts downstream of calcium mobilization in TFF2 driven repair

NHE2 has been previously implicated as acting downstream of TFF2 action, in an unknown manner, to promote gastric repair in vivo (Xue et al., 2011). To determine if

NHE2 was necessary to repair within the in vitro gastric organoid model and investigate if it affected Ca2+ mobilization, the selective NHE1/2 inhibitor Hoechst 694 (Hoe 694,

100µM) was pre-incubated in YC Nano gastric organoids prior to photodamage. At 10 min following damage Hoe 694 delayed epithelial repair, with a damage area of 32.03 ±

7.53 µm2 and repair rate of 0.28 ± 0.04 min-1 versus damage area of 3.50 ± 2 µm2 and repair rate of 0.49 ± 0.05 min-1 in control (Figure 4.10A, B) (both n=4, p<0.05).

Interestingly, Hoe694 did not significantly alter the Ca2+ mobilization following damage

(Figure 4.10C, D) (control 1.37 ± 0.04 vs Hoe 694 1.30 ± 0.02, both n=4). These results show that while NHE2 action is important for repair, it does not affect Ca2+ mobilization during the repair process, suggesting NHE2 acts downstream of Ca2+ mobilization during repair.

Consistent with these findings, as well as prior in vivo work (Xue et al., 2011),

NHE2 KO gastric organoids exhibited a significantly delayed exfoliation of 4.21 ± 0.66

µm compared to the WT gastric organoid exfoliation of 8.81 ± 0.70 µm (Figure 4.10E)

(n=5, p<0.05). To confirm that NHE2 acts downstream of TFF2 action during repair as suggested by previous in vivo studies (Xue et al., 2011), rTFF2 was microinjected into

NHE2 KO organoids and monitored over time.

Microinjection of rTFF2 into NHE2 KO organoids did not stimulate exfoliation (4.61 ± 0.41

µm, n=6, p<0.05) (Figure

4.10E). To confirm NHE2 is involved in TFF2 driven repair, TFF2 KO organoids were used to test whether

Hoe 694 would affect rTFF2 rescue action

(Figure 4.10F). Similar to earlier results measuring

Figure 4.10. Effect of NHE1/2 inhibition and loss of NHE2 function on calcium mobilization and repair. Fluorescence of YC Nano gastric organoids was imaged over time in panels A-D, and cell exfoliation was measured over time in panels E-F. Where indicated, Hoe 694 (100 µM) was added to organoid medium 1 hr prior to experimentation. In time courses, PD occurred at t=0 min. A) Damage area measured in YC Nano control (black) and Hoe 694 supplemented gastric organoids (blue) (n=4). B) Comparison of rate of repair between YC Nano control (black) and Hoe 694 supplemented gastric organoids (blue) (*p<0.05). C) Measurement of normalized FRET/CFP ratio of lateral membrane region of cells adjacent to the damage site comparing control (black) and Hoe 694 supplemented gastric organoids (blue). D) Comparison of the maximum FRET/CFP ratio from panel C between control (black) and Hoe 694supplemented gastric organoids (blue) (n=4, *p<0.05). E) Comparison of exfoliation in WT (n=5) and NHE2 KO vehicle (n=5) and rTFF2 injected organoids (n= 6) (*p<0.05). F) Comparison of exfoliation in WT and TFF2 KO gastric organoids treated with Hoe 694 and/or microinjection of rTFF2. Vehicle (WT Control, n= 5; TFF2 KO, n=6; TFF2 KO + rTFF2, n =4), Hoe 694 (WT, n=5; TFF2 KO, n=4; TFF2 KO + rTFF2, n=4). *p<0.05 vs WT vehicle, #p<0.05 vs rTFF2 treatment in TFF2 KO.

damage area (Fig 10A), WT control exfoliation (9.66 ± 1.05 µm, n=5) was significantly inhibited by addition of Hoe 694 (1.90 ± 0.56 µm, n=5, p<0.05). Reduced exfoliation was again observed in TFF2 KO (1.26 ± 0.32 µm, n=6) and addition of Hoe 694 did not inhibit exfoliation further (1.33 ± 0.24 µm, n=4). Microinjection of rTFF2 rescued exfoliation (9.0 ± 0.71 µm, n=4); however, the presence of Hoe 694 prevented the rescue effect of rTFF2 (1.42 ± 0.23 µm, n=4, p<0.05). These data further support NHE2 as necessary for the repair process and a likely downstream target of TFF2 action during repair.

4.4.8 Confirmation of efficacy of inhibitor concentrations used

To confirm the efficacy of the concentrations used, based upon previous in vitro studies (Yao et al., 2001; Chen et al., 2002; Aihara et al., 2018), experiments were performed evaluating AMD3100, AG1478 and Hoe 694 at difference concentrations

(Figure 4.11A-F). In the presence of 0.5 µM AMD3100, half the concentration used in the previous experiments presented, did not affect exfoliation or rate of repair compared to the control (Figure 4.11 A, B). However, at 1 µM and 3 µM AMD3100, both exfoliation and rate of repair were significantly decreased compared to the control

(Figure 4.11 A, B). While the concentration of 200 nM AG1478 has been demonstrated to be effective in inhibiting EGFR function (Chen et al., 2002), we wished to confirm that the efficacy of this concentration. While 200 nM and 300 nM AG1478 significantly delayed exfoliation and rate of repair compared to control, 100 nM AG1478 had no significant effect (Figure 4.11 C, D). In organoids treated with Hoe 694, only the concentration of 50 µM Hoe 694 did signifcantly affect exfoliation or rate of repair compared to control (Figure 4.11 E, F). These data suggest that the concentrations in

experiments shown previously are used at effective concentration. Furthermore, in all

cases, no additional effect

of increasing the inhibitor

concentration 3-fold above

the concentrations used,

suggesting the inhibitor

effects were saturable and

maximal effects were

observed.

Figure 4.11 Effect of inhibitor concentrations on exfoliation and repair. A) Time course of exfoliation in control (black, n=5) and concentrations of 0.5 µM (red circle, n=4), 1 µM (red triangle, n=5), and 3 µM (red square, n=5) AMD3100 supplemented gastric organoids following PD at t=0 min. B) Comparison of rate of repair between control (n=5) and concentrations of 0.5 µM (n=4), 1 µM (n=5), and 3 µM (n=5) AMD3100 supplemented gastric organoids. *p<0.05. C) Time course of exfoliation in control (black, n=4) and concentrations of 100 nM (green circle, n=7), 200 nM (green triangle, n=6), and 600 nM (green square, n=6) AG1478 supplemented gastric organoids following PD at t=0 min. D) Comparison of rate of repair between control (n=4) and concentrations of 100 nM (n=7), 200 nM (n=6), and 600 nM (n=6) AG1478 supplemented gastric organoids. *p<0.05. E) Time course of exfoliation in control (black, n=6) and concentrations of 50 µM (blue circle, n=3), 100 µM (blue triangle, n=4), and 300 µM (blue square, n=6) Hoe 694 supplemented gastric organoids following PD at t=0 min. F) Comparison of rate of repair between control (n=6) and concentrations of 50 µM (n=3), 100 µM (n=4), and 300 µM (n=6) Hoe 694 supplemented gastric organoids. *p<0.05.

4.5 Discussion

Restitution is the initiating event in epithelial repair and involves cell migration, not proliferation, for rapid re-establishment of epithelial integrity following injury. In this study we focused upon gastric epithelial restitution in response to microscopic photodamage, using a reductionist gastric epithelium model, gastric organoids. This model allows us to investigate the innate epithelial response separate from the complexities of native tissue, as the organoid system is devoid of other tissue cell types

(immune cells, mesenchymal cells, smooth muscle, neurons, etc.).

The current work provides deeper validation that the gastric organoid model maintains fidelity for major features of gastric restitution, as compared to in vivo photodamage results (Xue et al., 2010; Xue et al., 2011; Demitrack et al., 2012; Aihara et al., 2013; Aihara & Montrose, 2014). We have recently demonstrated that the gastric organoid system is comparable to native tissue in vivo in demonstrating the shedding of dead cells into the gastric lumen with an epithelial repair time course of ~10 min (Aihara et al., 2018). In the present study, gastric organoids are also found to be similar to native tissue as they demonstrate (1) increased intracellular Ca2+ mobilization during repair (Aihara et al., 2013), (2) dependence on TFF2, CXCR4, and NHE2 for repair

(Xue et al., 2010; Xue et al., 2011), and (3) placement of NHE2 as the most downstream effector identified in the TFF2/CXCR4 repair pathway (Xue et al., 2011).

These results identify features intrinsic to the epithelium, which operate in the absence of normal tissue architecture and accessory cell types. Furthermore this study delineates the relationship among known components of wound healing and links them within a signaling pathway, using an in vitro culture that more closely reflects native

tissue. Through the gastric organoid system we have been able to determine upstream and downstream effectors of gastric restitution which had been previously difficult to elucidate in vivo. This work now identifies specific pathways that can be tested in vivo in future work.

The gastric organoid system is reported to contain various cell types as seen in vivo, as well as exhibit similar responses to infection and damage as native tissue

(Schumacher et al., 2015a; Schumacher et al., 2015b; Aihara et al., 2018). Despite having a diversity of cell types within the organoid system, damage repair has not shown a heterogenous response either in this study or previous work (Aihara et al.,

2018). Different cell types were not morphologically identifiable, and no criteria were applied during the selection of cells to undergo damage, beyond their physical location in a site amenable to optical tracking of repair and cell extrusion. This observed homogeneity in the cellular response to damage suggests that this function is not dependent on cell type but is a highly conserved response to prevent loss of epithelial barrier function.

This is the first study to utilize the genetically encoded YC Nano Ca2+ reporter in studies of gastric tissues. Previously, our lab used YC 3.0 transgenic mice to show that endogenous Ca2+ mobilization is required for proper wound repair in vivo (Aihara et al.,

2013). The gastric organoid model, utilizing the more sensitive YC Nano Ca2+ indicator, offers a significant improvement over in vivo techniques as Ca2+ levels in individual cells can be resolved using a greater dynamic range of FRET/CFP ratio change, and a brighter overall signal (see Chapter 3). Using YC Nano gastric organoids, we show that intracellular Ca2+ mobilization is a downstream event stimulated by TFF2, CXCR4 and

EGFR activity during the repair process. Using the enhanced imaging resolution of organoids, we determined that Ca2+ mobilization was largely restricted to the cells directly adjacent to the wound site. Further, within these cells the lateral membrane region adjacent to damage was a proverbial hot spot of Ca2+ mobilization. Recently, we demonstrated that actin increases in the lateral membrane to initiate restitution and this action requires calcium and CXCR4 (Aihara et al., 2018). We speculate that this subcellular region may be a localized area optimized to stimulate Ca2+-dependent biochemical events, such as actin dynamics, in the part of the cell mediating cell motility. While Ca2+ mobilization is demonstrated to be important in repair, further studies are necessary to understand the source of this raised cytosolic Ca2+; whether it is the direct result of Ca2+ released from intracellular stores, and/or from activation of

Ca2+ flux across the plasma membrane.

Several lines of evidence support a link between TFF2 and Ca2+ mobilization. In

Jurkat cells, TFF2 activates Ca2+ signaling via the CXCR4 chemokine receptor

(Dubeykovskaya et al., 2009). In colonic Caco-2 epithelial cells, activation of CXCR4 stimulated the release of intracellular Ca2+ and enhanced intestinal epithelial restitution through reorganization of the actin cytoskeleton (Agle et al., 2010). It has also been reported that gastric epithelial damage is associated with intracellular and extracellular

Ca2+ mobilization in vivo and this flux of Ca2+ is required to mediate tissue repair (Aihara et al., 2013).

While it has previously been shown in separate studies (and distinct cell types) that TFF can promote Ca2+ mobilization or Ca2+ can modulate repair, our study is the first to provide direct evidence that causally links and extends these observations. We

show that Ca2+ mobilization is required for TFF2-mediated repair which occurs via

CXCR4. The key observations are that inhibition of CXCR4 impedes Ca2+ mobilization and slows repair in normal tissue, and CXCR4 is also required when exogenous rTFF2 is added to rescue repair in TFF2 KO cells. Interestingly, addition of exogenous rTFF2 either to the basolateral (via media) or apical (via microinjection) side of the organoids has a similar ability to rescue the delayed repair observed in TFF2 KO organoids. This suggests that TFF2 can act either apically or basolaterally at the cell membrane. Other studies in vivo have shown that intravenous administration of 125I-TFF2 can rapidly be distributed to the basolateral domain of gastric neck cells and parietal cells; over a longer time, the radioactivity of125I-TFF2 was observed at the luminal surface or mucus layer in the stomach (Poulsen et al., 1998). Furthermore, TFF has been detected in both serum and luminal fluid of the GI tract (Aikou et al., 2011). It is generally assumed that the route of TFF2 action via luminal secretion of TFF2 from the mucus cells lining the stomach; TFF2 has been shown to be secreted into the gastric lumen in parallel with mucus secretion (Kjellev et al., 2006). Our results support that TFF2 may be capable of acting at either the luminal or intravenous site; suggesting the presence of receptor binding sites at the cell membrane. Normal repair, rTFF2-rescued repair, and damage- induced Ca2+ mobilization can all be blocked by the Ca2+ chelator BAPTA. This demonstrates the calcium dependence of the repair process and provides strong evidence that Ca2+ mobilization is an essential downstream effector of TFF2/CXCR4 action during repair.

Evidence outside of the GI tract, as well as studies with GI cancer cell lines, suggest that CXCR4 and EGFR may act via the same repair pathway, introducing the

concept of CXCR4 activating EGFR during the repair process (Billadeau et al., 2006;

Guo et al., 2007; Cheng et al., 2017). There is also evidence suggesting the ability of

TFF2 to either directly or indirectly activates EGFR in colonic cancer cells during cell invasion (Rodrigues et al., 2003; Kosriwong et al., 2011). Further, ERK is an integration point for multiple receptor-mediated pathways. There is also evidence in vitro that TFF2 treatment causes activation of ERK1/2 through the CXCR4 receptor in gastric cancer epithelial AGS cells and lymphocytic cancer Jurak cells (Dubeykovskaya et al., 2009), suggesting that TFF2 activation of CXCR4 mediates ERK signaling. Studies in Caco2 cells show that ERK phosphorylation during repair is attenuated by EGFR inhibition, indicating that ERK phosphorylation is triggered via a pathway involving EGFR activation (Buffin-Meyer et al., 2007). Stimulation of EGFR and subsequent activation of

ERK1/2 have been demonstrated to be present in healing gut mucosa (Hansson et al.,

1990), although MEK/ERK signaling is not always essential for restitution (Frey et al.,

2004) possibly due to region- or tissue-specific effects. There is additional evidence that

ERK1/2 activation is primarily responsible for TFF mediated initiation of healing. Yu et al. found that TFF2 enhanced cell migration and wound healing in the gastric cell line

AGS and rat small intestine cell line IEC-6 in an ERK1/2 activation dependent manner

(Yu et al., 2010).

Our data suggest that EGFR potentially acts downstream of CXCR4 and as a necessary component during TFF2-driven, however further research will need to be done to determine whether this is by transactivation or if EGFR acts independently of

CXCR4. Further, our results indicate that ERK1/2 activity is a necessary component for proper repair in the epithelium, although it has not been formally addressed whether

phosphorylation of ERK1/2 in this cascade is the direct effect of either CXCR4 or EGFR activation. Our data show that ERK1/2 acts upstream of intracellular Ca2+ mobilization during the repair process. Evidence from previous studies and current literature suggest

ERK1/2 may be the primary pathway of EGFR action during repair. Future studies are needed to confirm whether ERK is acting in the same pathway as TFF2 (or EGFR) during repair in the gastric epithelium.

Previously, our lab has shown that in vivo, NHE2 is necessary during the repair process and likely acts downstream of TFF2 during repair (Xue et al., 2011). Studies described here have extended these findings, as addition of exogenous rTFF2 to NHE2

KO organoids did not alter delayed repair, and NHE1/2 inhibition slowed repair of normal organoids. EGF contribution to restitution has been shown to be mediated in part by stimulation of NHE in gastric epithelial cells (Yanaka et al., 2002). EGF is involved in acute regulation of cytoskeletal elements and NHE activity (Iwatsubo et al.,

1989; Ghishan et al., 1992; Furukawa & Okabe, 1997; Furukawa et al., 1999) and the

ERK pathway has also been shown as a critical component of NHE activation (Yoo et al., 2011; Muthusamy et al., 2012). We demonstrate that inhibition of NHE2 does not affect Ca2+ mobilization, suggesting that Ca2+ acts upstream of NHE2 or (less likely) that

NHE2 action is regulated via a parallel Ca2+-independent pathway. The role of NHE2 in promoting repair remains unknown, but based upon literature, we hypothesize that

NHE2 may regulate actin polymerization during repair and a necessary component for cytoskeletal structural rearrangements during migration (Vexler et al., 1996; Denker &

Barber, 2002).

Our work demonstrates that TFF2 acts via CXCR4 and EGFR signaling, including ERK activation, to drive Ca2+ mobilization and promote gastric repair. This work expands knowledge about the TFF2 signaling pathway (Dubeykovskaya et al.,

2009; Xue et al., 2010) and points to TFF2 and its activation of CXCR4 and EGFR as potential targets for promoting restitution. Additionally, these studies validate gastric organoids as a platform for studying repair and identifying potential future therapeutic targets.

Chapter 5

Sources of localized calcium mobilization during gastric organoid epithelial repair

Chapter 5 Sources of localized calcium mobilization during gastric organoid epithelial repair

5.1 Abstract

Using gastric organoids derived from transgenic mice expressing a fluorescent Ca2+ reporter (yellow cameleon-nano15; YC Nano), Chapter 4 shows that intracellular Ca2+ increases in cells directly adjacent to a damaged cell, and that this calcium rise is essential for prompt repair of damage. Using this Ca2+ sensor, we seek to use gastric organoids to investigate potential sources of this intracellular Ca2+ mobilization.

Photodamage (PD) and resultant cell death was induced to 1-2 gastric organoid epithelial cells by ~3 sec high intensity 840 nm light. As described in Chapter 4, change in intracellular Ca2+ was measured as FRET/CFP ratio, in cells adjacent to damaged cells. Inhibitors were used to test roles of Ca2+ channels (10 µM verapamil, 20 µM

YM58483), Phospholipase C (10 µM U73122), and IP3R (50 µM 2-APB). Inhibition of voltage gated Ca2+ channels (verapamil) or store operated calcium entry (YM58483) resulted in delayed repair and dampened intracellular Ca2+ response. Furthermore, inhibition of phospholipase C (U73122) or inositol trisphosphate receptor (2-APB) likewise resulted in delayed repair and dampened Ca2+ response. These results suggest both extracellular and intracellular Ca2+ sources are essential for supplying the

Ca2+ that stimulates repair using regulated signaling pathways. The findings implicate increased intracellular Ca2+ during repair is mediated via Ca2+ uptake via plasma membrane Ca2+ channels as well as intracellular Ca2+ release from the ER. Additionally, this work suggests that there are multiple sources to facilitate intracellular Ca2+ mobilization, and that each source is essential for proper repair. Collectively, this work

indicates the usefulness of YC Nano to further assess intracellular Ca2+ dynamics and further investigate the signaling cascade behind Ca2+-mediated repair.

5.2 Introduction

As a messenger, intracellular calcium (Ca2+) regulates a variety of cellular processes, including epithelial function and secretion in the stomach. While much of the mechanism during restitution remains unclear, calcium has been reported as one of its influential factors (Aihara et al., 2013; Aihara & Montrose, 2014; Aihara et al., 2018). It has recently been shown that both intracellular and extracellular Ca2+ are essential for efficient gastric epithelial restitution (Aihara et al., 2013). Furthermore, chelation of extracellular Ca2+ results in delayed repair within the gastric organoid model (Aihara et al., 2018). The proper repair of a disrupted epithelium is essential in the stomach to maintain a cohesive barrier of protection from the acidic luminal gastric contents. (Niv &

Banic, 2014).

While signaling in Ca2+ dynamics have been well studied in other systems much remains unclear within the context of gastric epithelial cells especially during repair. In intestinal epithelial cell lines, restitution has been shown to rely upon phospholipase C

(PLC) regulation of cytosolic Ca2+ and that inhibition of any PLC isoform results in repressed restitution (Rao et al., 2007). PLC is known to catalyze the hydrolysis of phospholipid phosphatidylinositol (4, 5)-bisphosphate to generate diacylglycerol (DAG) and inositol trisphosphate (IP3) which are demonstrated to regulate various cellular

2+ processes, including the role of IP3 in release of Ca from the endoplasmic reticulum

(ER) (Berridge, 1997; Khare et al., 1997; MA et al., 2003). In airway epithelium,

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2+ cytosolic Ca oscillation is induced by extracellular ATP which mediates PLC and IP3 signaling pathway to release Ca2+ into the cytosol from the ER (Evans & Sanderson,

1999).

The general role of Ca2+ as an effector of gastric wound repair has been established for 30 years (Miller & Henagan, 1979; Critchlow et al., 1985; Takeuchi et al.,

1985; Cheng et al., 2001), however the mechanisms by which Ca2+ mobilization affects restitution has yet to be elucidated and the source of this Ca2+ mobilization remains largely unknown.

The key role of Ca2+ in the stomach has been demonstrated by the use of

2+ inhibitors. Addition of PLC inhibitors, IP3 receptor antagonist, or voltage-gated Ca channel blockers slowed gastric repair while also blocking Ca2+ mobilization stimulated in the restituting gastric epithelial cells (Aihara et al., 2013), suggesting intracellular Ca2+ mobilization depends on both intracellular and extracellular sources. Activation of phospholipase C (PLC) is a known initiator of Ca2+-dependent signaling. Activated PLC cleaves the lipid phosphatidylinositol to release inositol triphosphate (IP3) and diacylglycerol (DAG), metabolites that can stimulate Ca2+ release from intracellular stores and activate protein kinase C (PKC). PKC is also a conventional target of Ca2+- dependent regulation. Both PLC and PKC have been shown in primary, immortalized, and cancer-derived cell lines to stimulate epithelial cell migration (Ranta-Knuuttila et al.,

2002; Rao et al., 2007; Saidak et al., 2009). In human gastric cancer (AGS) cells, Ca2+ mobilization has been shown to stimulate repair from aspirin and deoxycholate induced damage (Redlak et al., 2007, 2008). The Ca2+-dependent PLC-beta isoforms are known to be localized in gastric surface cells (Miller & Henagan, 1979; McGarrity et al., 1996).

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While these studies point to the overarching role of endogenous Ca2+ in gastric epithelial repair, due to systemic effects and difficulties in vivo little is known about the signaling needed to regulate and promote Ca2+ mobilization.

Due to the difficulty of studying gastric restitution and its signaling cascade in vivo, which is limited by suitable inhibitors as well as the tools to manipulate and monitor intracellular calcium, we sought to use the organoid culture system to investigate the signaling cascade behind Ca2+ mobilization during gastric epithelial repair and confirm previous in vivo findings. The gastric organoid culture system contains the cell types found in normal native tissue (Bartfeld et al., 2015; Bertaux-Skeirik et al., 2015;

Schumacher et al., 2015a) and allow for the growth and differentiation of epithelial cells.

We have recently acquired the Yellow Cameleon Nano15 (YC Nano) transgenic mouse, a more sensitive calcium sensor that utilizes FRET imaging to indicate changes in intracellular calcium. By using gastric organoids derived from YC Nano transgenic mice, we seek to investigate the potential sources of intracellular calcium mobilization during repair.

5.3 Materials and Methods

5.3.1 Animal husbandry

Experiments used transgenic mice (C57BL/6 background) expressing the Yellow

Cameleon-Nano15 (YC Nano) Ca2+ sensor fluorescent proteins (Oshima et al., 2014).

Pups were genotyped by genomic PCR as previously described (Schultheis et al., 1998;

Bell et al., 1999; Farrell et al., 2002) and used for experimentation at 2-4 months of age.

Animals were given standard rodent chow diet and water, both ad libitum. All animal

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procedures were approved by the Institutional Animal Care and Use Committee of the

University of Cincinnati.

5.3.2 Mouse-derived corpus organoid culture

Gastric organoids were generated from mouse gastric corpus as previously described (Mahe et al., 2013; Schumacher et al., 2015a; Engevik et al., 2018b). Isolated gastric epithelium from the corpus was cultured in Matrigel diluted 1:1 in Dulbecco’s

5.3.3 Induction of two-photon laser-induced photodamage

NIR 40x objective lens. To assess repair of damage area, images of YFP (excitation

514 nm, emission 535–590 nm) in the gastric organoid were collected simultaneously with transmitted light and a confocal reflectance image (reflecting 730 nm light to show cell/tissue structure). For assessing intracellular Ca2+ changes in YC Nano gastric organoids, images of YFP-FRET (Ti-Sa excitation 840 nm, emission 535-590 nm) and

CFP (Ti-Sa excitation 840 nm, emission 500-530 nm) were collected simultaneously with a transmitted light image. Wavelength selections for Ca2+ imaging were guided by

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previous work with YC sensors (Horikawa et al., 2010; Oshima et al., 2014). In all photodamage (PD) experiments, after collecting a set of control images, a small rectangle region (≈5 µm2) of a single cell was repetitively scanned at high Ti-Sa laser power (730 or 840 nm: 630 mW average) for 500 iterations (requiring ≈3 s).

Experiments examined gastric organoids embedded in Matrigel, located approximately 100-300 μm from the cover glass. Inhibitors were pre-incubated at least 1 hr prior to experimentation to assure equilibration in Matrigel and were kept in the medium during experiments. Inhibitory reagents included: Verapamil (10 μM, Sigma),

YM58483 (20 μM, Tocris), U73122 (10 μM, Cayman Chemical), and 2-APB (50 μM,

Tocris). Final DMSO concentration in experiments was <0.1%. Solvent control groups contained 0.1 % DMSO added to medium. Vehicle control groups contained either 0.1%

DMSO, ddH2O, or dPBS added to the medium; vehicle was dependent on the solution the inhibitors used were constituted in. Concentrations were determined based upon in vitro studies and tested to assess effectiveness and potential toxicity.

Damage-repair cycle was measured independently once per gastric organoid, and outcomes from at least 3 different gastric organoids (derived from at least 3 animals), were compiled for each experimental protocol.

5.3.4 Image analysis

PA, USA). The damaged area was measured as the region of cellular loss of YFP

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fluorescence in YC Nano gastric organoids. In each experiment of YC Nano gastric organoids, we determined the time point displaying maximal damage area and estimated rates of epithelial restitution starting from this time with a single exponential curve fit to the size of damage area over time (Xue et al., 2010; Aihara et al., 2018).

Best fit values of the rate constant were used as estimates of the rate of repair (units of min−1). Changes in intracellular Ca2+ were measured as FRET/CFP ratio using YC Nano gastric organoids. Background images were subtracted from FRET-YFP and CFP images, the resultant images were divided on a pixel-by-pixel basis to calculate the

FRET/CFP ratio image. All time course ratio images were then normalized to the averaged pre-damage baseline images. Regions of interest (ROIs) were determined by transmitted light and 514 nm excited YFP images to define cellular structures for whole cell and lateral region measurements.

5.3.5 Bioinformatics RNA sequencing data analysis

RNA sequencing data was acquired from public repository (GEO accession number: GSE73336) (Engevik et al., 2016). The differential gene expression analysis was performed based on the negative-binomial statistical model of read counts as implemented in the Bioconductor packages DESeq (organoid vs fundic tissue samples)

(Anders et al., 2013). Significance tests are not considered for the organoid vs fundic tissue samples because the groups consist of single replicates.

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5.3.6 Statistical analysis

5.4 Results

5.4.1 Identifying potential calcium targets in gastric organoids

To identify potential Ca2+ targets present in both native tissue and gastric organoids, we utilized previously published RNA sequencing (Engevik et al., 2016) and generated a list of 166 gene targets whose annotations indicated they were involved in

Ca2+ signaling (Figure 5.1, Table 5.1). It should be noted that in the RNA seq list, duplicates of 3 genes appear because of multiple sequences being separately evaluated within those genes.

Out of the 166 targets, there was notable concordance between the expression in organoids and tissue.

Among those 46 targets that were most highly expressed in tissue (expression higher than 150 reads), 35 of those targets were also the most highly Figure 5.1 Heatmap of potential targets involved in calcium signaling within gastric organoids and tissue, based upon expressed in organoids. Notably, RNA sequence data set acquired from Engevik AC, et al., CMGH, 2016 GSE73336.

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among those genes were inositol trisphosphate receptor (IP3R) type 3 (itpr3), Calcium

Voltage-Gated Channel Subunit Alpha1 D (cacna1d), Calcium Voltage-Gated Channel

Auxiliary Subunit Beta 3 (cacnb3), Calmodulin 1 (calm1), and Calmodulin2 (calm2).

Among the targets expressed at the lowest levels in tissue and organoid (expression below a cut-off of 20 reads), 105 targets were the lowest in organoids and 78 targets were at lowest levels in tissue. Notably, the ryanodine receptor type 1 (ryr1) was among this set, and all other ryanodine receptors (ryr2, ryr3) had expression below 68 reads in both tissue and organoids. It should be noted that there were Ca2+ targets that were highly expressed in one sample but at low or nil expression in the other, demonstrating that the organoids were not a perfect mimetic of native tissue. However, results suggest that calcium handling in organoids can be expected to be broadly similar between organoids and tissue.

Among the highest Table 5.2 Table of calcium targets with the highest RNA expression in both tissue and organoid based upon Table 5.1 expression found in both organoid and tissue sample, we chose to target voltage gated Ca2+ channels and inositol trisphosphate (IP3R)

(Table 5.2). In addition to these targets, based upon previous in vivo and in vitro work, store operated Ca2+ entry (SOCE) and phospholipase C were also chosen to examine Ca2+ signaling.

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Table 5.1 Table of RNA expression of 166 genes of interest based upon RNA sequence data.

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5.4.2 Voltage gated calcium channels are essential for intracellular calcium mobilization during repair

As shown in vivo using an older version of the YC cameleon (YC 3.0) Ca2+ indicator, blocking voltage gated Ca2+ channels via verapamil delays repair as well as dampens the intracellular calcium response (Niv & Banic, 2014). Our experiments in organoids used the YC Nano sensor to measure intracellular Ca2+ changes within intact cells neighboring the damage site via ratiometric imaging (Methods), and these experiments had the goal to confirm the necessity of voltage gated Ca2+ channels during repair and that the gastric organoid can recapitulate in vivo events. To verify the importance of voltage gated

Ca2+ channels during repair, verapamil was applied 1 hr prior to photodamage (PD). Figure 5.2 Effect of voltage gated calcium channel blocker on calcium mobilization and repair. Fluorescence of YC Nano gastric organoids was imaged over time. Verapamil (10 µM) was added to organoid medium 1 hr prior to experimentation. In time course, PD Following PD, the occurred at t=0 min. A) Measurement of normalized FRET/CFP ratio of lateral membrane region of cells adjacent to the damage site comparing control (black) and verapamil supplemented gastric organoids (green). Comparison of the maximum FRET/CFP ratio presence of verapamil from panel B between control (black, n=3) and verapamil treated (green, n=5) gastric organoids (*p<0.05). C) Comparison of rate of repair control (black, n=3) and verapamil dampened Ca2+ treated (green, n=5) gastric organoids (*p<0.05). mobilization compared to the control (Figure 5.2A), significantly blunting Ca2+ signaling within cells adjacent to the damage site, where the control FRET/CFP ratio peak was

1.32 ± 0.03 compared to a FRET/CFP ratio peak of 1.19 ± 0.02 (n=3) in the presence of verapamil ( n=5, p<0.05) (Figure 5.2B). Furthermore, the presence of verapamil caused

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a significantly delayed repair rate of 0.27 ± 0.08 min-1 (n=5) compared to the control repair rate of 0.59 ± 0.08 min-1 (n=3, p<0.05) (Figure 5.2C). These data suggest the contribution of voltage gated Ca2+ channels in the intracellular Ca2+ response during repair. Results support prior results in native tissue, and further confirm the use of gastric organoids as a valid model to elucidate Ca2+ signaling pathways during gastric repair.

5.4.3 Phospholipase C pathway is necessary for intracellular calcium mobilization during repair

We have previously shown in vivo that inhibition of PLC results in delayed repair and dampened calcium mobilization (Aihara et al., 2013), we have also shown in vitro that PLC inhibition affects actin mobilization as well as delays repair in gastric organoids

(Aihara et al., 2018).

To confirm this delay

observed in gastric

organoids is related to

Ca2+ mobilization, we

inhibited PLC by

Figure 5.3 Effect of Phospholipase C inhibition on calcium mobilization and repair. adding U73122 (10 Fluorescence of YC Nano gastric organoids was imaged over time. U73122 (10 µM) was added to organoid medium 1 hr prior to experimentation. In time course, PD occurred at μM) into the media t=0 min. A) Measurement of normalized FRET/CFP ratio of lateral membrane region of cells adjacent to the damage site comparing control (black) and U73122 supplemented gastric organoids (red). Comparison of the maximum FRET/CFP ratio from panel B and monitored the between control (black, n=4) and U73122 (red, n=6) gastric organoids (*p<0.05). C) Comparison of rate of repair control (black, n=4) and U73122 (red, n=6) gastric organoids response following (*p<0.05). injury. Following PD, U73122 dampened the Ca2+ response during repair (Figure 5.3A)

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which resulted in a significantly decreased FRET/CFP ratio of 1.15 ± 0.02 (n=6) compared to the control of 1.32 ± 0.03 (n=4, p<0.05) (Figure 5.3B). Furthermore, similar to what was observed in GFP-actin organoids (Aihara et al., 2018), U73122 significantly delayed the rate of repair (0.24 ± 0.01 min-1, n=6) compared to control

(0.56 ± 0.11 min-1, n=4, p<0.05) (Figure 5.3C). These data confirm previous findings in vivo and in vitro (Aihara et al., 2013; Aihara et al., 2018), identifying PLC as an important player in Ca2+ mobilization during epithelial repair.

PLC is reported to act via a signaling cascade to release of Ca2+ from the endoplasmic reticulum via the downstream receptor inositol triphosphate receptor (IP3R)

(Sambrook, 1990; Putney Jr & Ribeiro, 2000). Furthermore, according to RNA seq data,

IP3R was highly expressed in both fundic tissue and organoid sample suggesting IP3R as a potential target for

Ca2+ signaling. To Figure 5.4 Effect of inositol trisphosphate receptor inhibition on calcium mobilization and repair. Fluorescence of YC Nano gastric organoids was imaged over time. 2-APB (50 µM) was added to organoid medium 1 hr prior to experimentation. In time course, PD occurred at further confirm t=0 min. A) Measurement of normalized FRET/CFP ratio of lateral membrane region of cells adjacent to the damage site comparing control (black) and 2-APB supplemented gastric PLC’s role and organoids (purple). Comparison of the maximum FRET/CFP ratio from panel B between control (black, n=3) and 2-APB (purple, n=4) gastric organoids (*p<0.05). C) Comparison of rate of repair control (black, n=3) and 2-APB (purple, n=4) gastric organoids (*p<0.05). determine whether

the ER release of calcium is contributing to repair, we investigated the role of IP3R.

Gastric organoids were treated with an IP3R inhibitor (2-APB, 50 μM) to test the role of

2+ IP3R during restitution. Following PD, 2-APB dampened the Ca mobilization (Figure

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5.4A), resulting in a significantly decreased maximum FRET/CFP ratio of 1.26 ± 0.02

(n=4) compared to the control (1.67 ± 0.01, n=3, P<0.05) (Figure 5.4B). Furthermore, addition of 2-APB resulted in a significantly delayed repair rate (0.28 ± 0.03, n=4) compared to the control (0.68 ± 0.12, n=3, p<0.05) (Figure 5.4C). These data suggest

2+ the role of ER-released Ca via IP3R as a contributing source to promote epithelial repair.

5.4.4 Store operated calcium entry is essential for calcium mobilization during restitution

To further confirm extracellular Ca2+ is an essential source to aid in Ca2+ mobilization during gastric repair, an inhibitor of store operated Ca2+ entry (SOCE,

YM58483, 20 μM) was added to the media 1 hr prior to PD. The presence of YM58483

dampened the Ca2+

mobilization following

PD (Figure 5.5A),

resulting in a

significantly

Figure 5.5 Effect of store operated calcium entry inhibition on calcium mobilization and decreased repair. Fluorescence of YC Nano gastric organoids was imaged over time. YM58483 (20 µM) was added to organoid medium 1 hr prior to experimentation. In time course, PD occurred at maximum t=0 min. A) Measurement of normalized FRET/CFP ratio of lateral membrane region of cells adjacent to the damage site comparing control (black) and YM58483 supplemented gastric organoids (orange). Comparison of the maximum FRET/CFP ratio from panel B between FRET/CFP ratio at control (black, n=3) and YM58483 treated (orange, n=7) gastric organoids (*p<0.05). C) Comparison of rate of repair control (black, n=3) and YM58483 treated (orange, n=7) gastric organoids (*p<0.05). 1.2 ± 0.01 (n=7) compared to the control FRET/CFP peak at 1.37 ± 0.02 (n=3, p<0.05) (Figure 5.5B).

Additionally, inhibition of SOCE significantly delayed the rate of repair, reducing from a control 0.68 ± 0.13 min-1 (n=3, p<0.05) to 0.33 ± 0.06 min-1 (n=7) (Figure 5.5C). These

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data further suggest the role of extracellular Ca2+ contributing to the intracellular calcium response during repair.

5.5 Discussion

Ca2+ signaling is a dynamic process involving changes in intracellular Ca2+ availability as well as coordination of Ca2+ release from surrounding cells following epithelial damage (Sanderson et al., 1994). As a second messenger, Ca2+ is capable of acting in several different signaling cascades each of which has a different mechanism by which Ca2+ mobilizes. Some of such mechanisms have been shown to cause Ca2+ mobilization that is highly localized, brief increases of Ca2+ while other pathway mechanisms produce longer-lasting elevations of Ca2+ which often follow oscillations caused by feedback loops within the signaling system (Berridge, 1997). In vivo gastric damage elicits increased gastric luminal Ca2+ (Takeuchi et al., 1985; Koo, 1994;

Takeuchi et al., 1999). Intracellular Ca2+ has also been shown to be essential for proper gastric wound repair (Aihara et al., 2013). In vitro inhibition of PKC or Ca2+ mobilization significantly inhibits cell migration following wounding in cultured rabbit gastric cells

(Ranta-Knuuttila et al., 2002). While intracellular Ca2+ appears to be localized within the cells adjacent to the damage site, studies have been limited in investigating Ca2+ mobilization within an in vitro model that closely mirrors in vivo native, non-cancerous tissue. Work from the Hyser lab has demonstrated (using the GCamp6s Ca2+ sensor in human jejunum enteroids) that rotovirus infection activates dynamic Ca2+ signaling through mediation of SOCE and purinergic signaling in infected cells (Chang-Graham et al., 2018). This speaks to the ability of viruses to utilize innate cellular signaling to

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induce pathophysiological signaling, while also displaying dynamic Ca2+ signaling among cells under normal conditions.

We have previously shown in vivo the importance of both extracellular and intracellular Ca2+ during gastric epithelial repair (Aihara et al., 2013; Aihara & Montrose,

2014). This study was the first to demonstrate the Ca2+ dynamics occurring during gastric restitution with a genetically encoded Ca2+ indicator (GECI) , however due to tissue motion in a breathing animal it was limited in its ability to observe Ca2+ dynamics in individual cells during the response to single cell damage. In vivo models are also limited in the ability to apply drugs that do not have more systemic effects. Similar to what was observed in vivo, the current study found that inhibition of voltage gated

2+ channels, PLC and IP3R, caused significantly decreased Ca mobilization in addition to delayed restitution. Our findings parallel what has been witnessed in vivo, indicating the reliability of gastric organoids to mimic native tissue and confirming the role of both voltage gated channels and the PLC pathway during Ca2+ driven repair.

One known pathway responsible for releasing intracellular Ca2+ stores from the endoplasmic reticulum (ER) involves PLC. Either in response to activation via a G- protein coupled receptor or tyrosine kinase receptor, PLC hydrolyzes the IP3 precursor, phosphatidylinositol 4,5-bisphosphate (PIP2) to produce diacylglycerol (DAG) and IP3

(Berridge, 1993). Following hydrolysis of PIP2, IP3 binds to its receptor IP3R which undergoes a conformational change that leads to the mobilization of stored Ca2+ from the ER (Mignery & Südhof, 1990). IP3R, along with ryanodine receptors (RYRs), are the principal intracellular Ca2+ channels responsible for the release of Ca2+ from the ER membrane stores (Berridge, 1993). Based upon RNA sequencing (Table 1), we chose

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2+ to test IP3R as an indicator of ER release of Ca stores due to its high expression of

IP3R Type 3 and Type 1 in both gastric organoids and tissue. While the RYR family was a potential candidate, the RNA sequencing data showed that all RYR family members had negligible to nil expression within either gastric organoids or intact tissue. Our data

2+ shows that IP3R is a necessary component to facilitate intracellular Ca mobilization during repair, indicating that the ER plays an important role in supplying Ca2+. As SOCE is reported to be activated by release of ER Ca2+ stores (Parekh & Penner, 1997;

2+ Parekh, 2003), it is possible that SOCE acts following the release of Ca via IP3R.

However further studies are needed to determine the order within the signaling cascade.

This study expands our current understanding of Ca2+ dynamics by demonstrating the importance of two classes of plasma membrane channels and the role of intracellular

(ER) release in the observed intracellular Ca2+ mobilization (Figure Figure 5.6 Schematic diagram of calcium mobilization during 5.6). These results offer evidence to epithelial restitution. Dotted lines indicate more speculative pathways. Ca2+, calcium; SOCE, store operated calcium entry;

PLC, phospholipase C; IP3, inositol trisphosphate; IP3R, inositol build future studies upon to further trisphosphate receptor. delve into the signaling cascade behind gastric epithelial restitution, with the ultimate goal of identifying druggable targets to improve gastric repair in compromised situations.

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Chapter 6

Discussion

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Chapter 6 Discussion

6.1 Significance of Outcomes

Collectively, our data provide new insight into gastric restitution. Addressing the precise epithelial signaling cascade associated with restitution in vivo is difficult due to the presence of immune cells, mesenchyme and other factors. Additionally, only a limited number of inhibitors and agonists are available for in vivo studies. Most prior in vitro work was based on cancer-derived gastric models which don’t reliably recapitulate native tissue. Our work with gastric organoids has offered a valuable new tool for examining gastric repair. This work has now established the two-photon induced damage model and Ca2+ mobilization in organoids; a tool that will overall add value to organoids in general for studying epithelial physiology. Our model provides a unique window into understanding the early factors in pathogenesis, which have the potential to lead to preventative interventions to block detrimental effects prior to tissue degradation.

By examining TFF2-, EGFR-, NHE2- and Ca2+-mediated restitution, we have drawn together separate fields and connected a number of findings that have previously existed in isolation. Prior to this work, no proposed models linked TFF2, EGFR, NHE2 and Ca2+ mobilization. This research is therefore conceptually innovative and provides the first evidence for TFF2 mediated EGFR signaling cascades driving NHE2 activity.

The translational relevance of these studies underscore the critical role of understanding TFF mediated gastric restitution. This work extends our current understanding of the role of TFF2 and NHE2 during gastric restitution. We have also identified the initiating events leading the Ca2+ mobilization and identified novel pathways by which Ca2+ may be entering the cells. We believe that this work has

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identified several key targets for restitution and in the future these targets may be used to promote gastric repair in patients.

Wound healing is a dynamic and complex process required to minimize damage to host tissue. The healing process is divided into four distinct phases: (1) restitution, (2) inflammation, (3) proliferation, and (4) maturation (Velnar et al., 2009). Restitution as the initiating phase of repair represents a critical step in the healing process and factors that promote or enhance restitution may be key in promoting downstream healing.

Multiple studies have demonstrated that supplementation of factors know to be involved in restitution, such as TFF, increase the overall wound repair (Playford et al., 1995;

Babyatsky et al., 1996; McKenzie et al., 1997; McKenzie et al., 2000; Farrell et al.,

2002; Iizuka & Konno, 2011; Shi et al., 2014; Leoni et al., 2015). Likewise, loss of select restitution promoting factors has been shown to slow wound repair (Mashimo et al.,

1996; Podolsky, 1999; Furuta et al., 2001; Beck et al., 2004; Podolsky et al., 2009); supporting the important role of restitution in general healing. Restitution also serves as an essential step in dead cell shedding and maintaining the epithelial barrier integrity.

The gastric epithelium is constantly exposed to exogenous (alcohol, aspirin, pathogen Helicobacter pylori) as well as endogenous (luminal acid, digestive enzymes) damaging agents (Podolsky, 1999; Taupin & Podolsky, 2003). Factors, including nonsteroidal anti-inflammatory drugs (NSAIDs) and H. pylori, slow epithelial repair which can lead to peptic ulcers (Kusters et al., 2006). Studies indicate that NSAID treatment slows the production and alters the structure of gastric mucus, reduces the hydrophobicity of the gastric mucosal surface, depletes prostaglandins, increases cell shedding, decreases cell proliferation at the ulcer margin region, reduces gastric

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mucosal blood flow and interferes with the repair of superficial injury (Baumgartner et al., 1986; Becker et al., 2004; Matsui et al., 2011). By interfering with mucus-secreted proteins such as TFF2, and the proper expulsion of cellular debris, NSAIDs can cause gastric injury by directly interfering with restitution (Wallace et al., 1990; Wallace, 2000).

Interference with the process of restitution in animal models has been shown to contribute to the conversion of superficial injury to deeper mucosal necrosis (Wallace et al., 1990), highlighting the importance of restitution in ulcer formation.

H. pylori infection is known to promote ulcers through multiple mechanisms. H. pylori has been reported to injure gastric epithelial cells by producing ammonia

(Mégraud et al., 1992), cytotoxins called CagA (cytotoxin-associated gene A) product

(Covacci et al., 1993) and Vac A gene product (vacuolating cytotoxin) (Cover & Blaser,

1992). Ammonia concentrations designed to match what is observed in H. pylori infected patients has been shown in vitro to impair gastric restitution (Yanaka et al.,

1993; Suzuki et al., 2000). In separate studies, H. pylori virulence factors VacA and

CagA exert no effect on epithelial cell migration or overall restitution (Ricci et al., 1996).

However, undialyzed H. pylori broth culture filtrates inhibited both cell migration and restitution through a VacA- and CagA-independent mechanism, indicating that H. pylori may secrete factors that specifically inhibit gastric restitution (Ricci et al., 1996).

Moreover, H. pylori infection has been found to promote methylation and silencing of

TFF2 (Peterson et al., 2010), which may further contribute to defects in repair.

Eradication of H. pylori has been shown restore gastric structure and repair (Kuipers et al., 2004; Van Grieken et al., 2004), providing additional evidence for the role of H. pylori in inhibition gastric restitution and global wound repair.

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Both NAIDS and H. pylori infection contribute to the development of Peptic ulcer disease (PUD). PUD affects more than 6 million people each year in the United States and continues to be a burden in medical care (Feinstein et al., 2010). Statistics from the

NIH report that PUD cost the U.S. healthcare system 1.9 billion dollars in 2004. Despite decreases in PUD incidence over the past decades, PUD still remains the lead diagnosis on 37% of hospital discharges (Everhart & Ruhl, 2009). Furthermore, new research suggests that proton pump inhibitors (PPIs), the most widely used treatment regimens for PUD, may increase the risk of dementia (Gomm et al., 2016) and chronic kidney disease (Moledina & Perazella, 2016; Xie et al., 2016). These latest findings highlight the need to better understand of the mechanisms of gastric restitution to help minimize ulcer incidence or accelerate the repair process.

The goal of this project was to understand the role of Ca2+ signaling in gastric restitution. This represents an important gastric defense mechanism that is vital to protect the underlying epithelium and initiate repair. Ca2+ ions partake in signaling networks that underlie the regulation of diverse cellular processes such as cellular communication, metabolism, transcription, protein folding, cellular proliferation, and actin rearrangement (Clapham, 2007). As a result, Ca2+ mobilization and distribution within cells is tightly regulated, and the disruption of Ca2+ homeostasis has been shown to be involved in multiple injury and disease states (Carafoli & Brini, 2007). In addition to

Ca2+, this project also sought to identify key factors involved in initiation of restitution including trefoil factor 2 (TFF2) and the sodium hydrogen exchanger isoform 2 (NHE2).

Utilizing a model of gastric damage caused by two-photon photodamage, this work is the first to detail how Ca2+ signaling, EGFR, and NHE2 are integral to mediating TFF2

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driven restitution. Although several groups have recognized the potential for TFF,

CXCR4, EGFR and Ca2+ to participate in restitution, the lack of physiologically relevant models in vitro have made it challenging to define the signaling cascade associated with restitution (Mori et al., 1987; Iwatsubo et al., 1989; Hansson et al., 1990; Wright et al.,

1990; Ménard & Pothier, 1991; Ghishan et al., 1992; Furukawa & Okabe, 1997;

Kinoshita et al., 2000; Chen et al., 2001; Yanaka et al., 2002; Rodrigues et al., 2003;

Dubeykovskaya et al., 2009; Xue et al., 2010; Yu et al., 2010; Xu et al., 2013). Our work utilizes the novel gastric organoid model, which closely resembles native tissue. Based on our work with gastric organoids, we demonstrate that TFF2 activates CXCR4 which drives Ca2+ signaling. We also demonstrate that the effect of EGFR is mediated through intracellular Ca2+ signaling. Furthermore, our results suggest EGFR as a potential downstream effector of TFF2 mediated repair, as both are essential aspects for proper repair. Interestingly, the effects of NHE2 do not impact Ca2+ signaling, indicating that

NHE2 activity is downstream of Ca2+. Our work points to intracellular Ca2+ at the lateral membrane region as a sensitive detector of mobilization following damage. This research is technically innovative because it utilized the use of two-photon microscopy to induce focal and controlled photodamage as well as monitored responses to challenges that occur in minutes, rather than days or weeks. Our lab has previously utilized photodamage in vivo (Demitrack et al., 2010; Xue et al., 2010; Xue et al., 2011;

Aihara et al., 2013) and in vitro (Aihara et al., 2018) to induce localized damage to 1-3 cells and monitor repair; unlike other damage models, photodamage allows for live imaging in real time. This work has now established methods for using the two-photon laser damage model and Ca2+ mobilization in organoids; tools that we believe will

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overall add value to organoids in general for studying epithelial physiology.

Furthermore, this work has shown that photodamage induces caspase-activated apoptosis which mirrors what occurs during natural cell shedding in organoids. This model provides a unique window into understanding the early factors in pathogenesis which results from a compromised epithelial layer, offering the potential to identify interventions to block detrimental effects prior to tissue degradation.

Diving deeper into the signaling cascade, we found that intracellular Ca2+

2+ originated from PLC and IP3R driven Ca release from the endoplasmic reticulum (ER).

This is consistent with other studies which demonstrate that the ER plays a prominent role in the regulation of cellular Ca2+ homeostasis (Paschen, 2001). Moreover, extracellular Ca2+ also acts as a source for Ca2+ mobilization via voltage gated calcium channels and Store Operated Calcium Channels (SOCE). Both sources of Ca2+ are tightly regulated and coordinated with one another, as Ca2+ release from the ER triggers the activation of plasma membrane Ca2+ channels in a process known as store- operated Ca2+ entry (SOCE) (Putney Jr, 2007). Our work extends the existing literature by definitively identifying intracellular and extracellular Ca2+ sources as vital for gastric restitution in organoids, which mimics prior conclusions in vivo. This work is among the first to then delineate the signaling cascades associated with Ca2+ -driven gastric restitution, however further studies are needed to assess the order by which these Ca2+ sources play a role. The work herein greatly improves our understanding of restitution associated intracellular signaling and may provide new targets for initiating wound healing. We have defined the pathway behind TFF2 signaling during gastric epithelial repair and linked pathways which previously existed in isolation. Our work has

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connected TFF2 activation of CXCR4 with activation of EGFR and activation of ERK1/2 and downstream Ca2+ mobilization. To the best of our knowledge, this is the first study that has examined these signaling cascades in a single study and the first to identify the co-requirement of these signaling factors in gastric restitution. This innovative examination of epithelial restitution may have a positive impact on our ability to improve therapeutics for promoting ulcer repair.

6.2 Future Directions

Our work has established the novel roles of TFF2 and its receptor, CXCR4,

EGFR, Ca2+ and NHE2 in gastric restitution within a valuable new model that offers a strong homology to cells of native tissue. The physiologically relevant organoid model has allowed us to examine these specific signaling cascades in greater depth than previous studies. To the best of our knowledge, our inhibitors are highly specific, and the concentrations have been previously used in literature (Chen et al., 2002; Hurst et al., 2008; Aihara et al., 2018) and tested in our organoid model for effects on viability and for their concentration dependency. However, it is possible that some of our inhibitors may have unforeseen off target effects. To circumvent this possibility, in the future organoids could be generated from knockout mice including CXCR4 (Odemis et al., 2005), L-type calcium channels (Moosmang et al., 2005), Orai1 (Gwack et al., 2008) and STIM1 (Mancarella et al., 2013). Use of these organoids could help address the precise signaling pathways involved in gastric restitution without the requirement of inhibitors and could simultaneously offer a platform for testing drug specificity. However, there is always the potential that congenital loss of these proteins could result in over-

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compensation of related transporters that would confound results. Recently groups have generated tamoxifen-inducible organoids from mouse models and have induced knock out the proteins of interest in organoids (Engevik et al., 2018a). This same approach could be used in our system. Additionally, activators of these target could also be used to confirm the importance of these systems for gastric repair.

In contrast to removing proteins of interest, an alternative that could be utilized is to over-express select proteins. For example, Vector Biolabs has mouse TFF2 adenovirus commercially available and this could be used to generate an overexpression model in mouse gastric organoids. This could also be used as a rescue experiment for TFF2 knockout organoids. To examine the calcium-related signaling cascades in greater depth, fluorescently tagged calcium-signaling proteins could be used. For example, GFP-tagged Orai1 and STIM1, the main determinants of the store- operated Ca2+ entry (SOCE), as well as L-type calcium channels have been previously used in cell culture (Zhang et al., 2005; Gwozdz et al., 2012) could be utilized to further examine Ca2+ dynamics, especially in the context of non-damage, and confirm our findings. These same approaches coupled with our live imaging and photodamage could be used in gastric organoids. These proposed studies would help extend our understanding of intracellular and extracellular Ca2+ signaling.

This research points to the role of TFF2 in mediating gastric restitution. Our experiments demonstrate that recombinant TFF2 can enhance restitution in organoids.

As a result, endogenous TFF2 may serve as potential therapeutic for conditions involved aberrant repair, including PUD. Recently, several groups have begun to engineer beneficial microbes, also known as probiotics, to secrete clinically prevalent

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compounds. For example, Lactococcus lactis sAGX0085 has been engineered to produce biologically active murine TFF in suitable quantities to allow for oral application of the L. lactis (Neirynck & Steidler, 2006). Daily intragastric administration of the L. lactis secreting murine TFF1 prior to or during dextran sodium sulfate colitis was shown to reduce mortality, weight loss in mice and improve colon histology compared to media broth control and L. lactis controls that do not secrete TFF1 (Neirynck & Steidler, 2006).

L. lactis has recently been developed to secrete human TFF1 and is currently being tested in a Phase 2 clinical trial for patients undergoing chemoradiation for the treatment of head and neck cancer to protect against oral mucositis (ClinicalTrials.gov Identifier:

NCT03234465) (Vargason & Anselmo, 2018). Although this approach has not yet been used in the stomach, L. lactis is an acid tolerant bacterium and has been shown in humans to a have 90% survival rate in the stomach (Drouault et al., 1999). As a result, it may be possible to use this approach in the future to deliver TFF2 for the treatment of gastric ulcers.

In patients, a number of disorders have been associated with decreased TFF2 diurnal rhythm. These disorders include H. pylori infection, aging, and sleep deprivation

(Hoffmann, 2005). These conditions are also all risk factors contributing to gastric ulcer formation. It has been demonstrated that H pylori-infected TFF2-deficient mice develop more advanced premalignant lesions than wild-type littermates (Fox et al., 2007).

Moreover, H. pylori infection has been shown to promote methylation and silencing of

TFF2 in mice and humans (Peterson et al., 2010). To confirm the effects of H. pylori on

TFF2 and repair, our novel gastric organoid two-photon damage model could be implemented. This work would definitely address whether H. pylori can silence TFF2

125

and delay restitution. Interestingly, in Mongolian gerbils a combination of a high salt diet with H. pylori significantly down-regulated TFF2 expression compared to H. pylori infection alone (Cheng et al., 2001). According to the Center for Disease Control (CDC),

90% of Americans consume more sodium than the daily recommendation of 2,300 mg/day (Jackson et al., 2016). To address the role of diet and infection, organoids could be microinjected with H. pylori in organoid media containing evaluated NaCl levels and restitution examined using our existing system. This work could result in novel findings for gastric restitution and overall for gastric wound repair.

126

Chapter 7

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