Functional Restoration of Irradiated Salivary Glands Through Modulation of Apkcζ and Nuclear Yap in Salivary Progenitors

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Functional Restoration of Irradiated Salivary Glands Through Modulation of aPKCζ and Nuclear Yap in Salivary Progenitors

Item Type text; Electronic Dissertation

Authors Martinez Chibly, Agustin Alejandro

Publisher The University of Arizona.

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Link to Item http://hdl.handle.net/10150/621771 FUNCTIONAL RESTORATION OF IRRADIATED SALIVARY GLANDS THROUGH MODULATION OF APKC AND NUCLEAR YAP IN SALIVARY PROGENITORS

Agustin Alejandro Martinez Chibly

A Dissertation Submitted to the Faculty of the

THE GRADUATE INTERDISCIPLINARY PROGRAM IN CANCER BIOLOGY

In Partial Fulfillment of the Requirements For the Degree of

DOCTOR OF PHILOSOPHY

In the Graduate College

THE UNIVERSITY OF ARIZONA

2016 2

THE UNIVERSITY OF ARIZONA

GRADUATE COLLEGE

As members of the Dissertation Committee, we certify that we have read the dissertation prepared by Agustin Alejandro Martinez Chibly entitled “Functional Restoration of Irradiated Salivary Glands Through Modulation of aPKC and nuclear Yap in Salivary Progenitors” and recommend that it be accepted as fulfilling the dissertation requirement for the Degree of Doctor of Philosophy

______Date: 09/08/16 Kirsten Limesand

______Date: 09/08/16 Jean Wilson

______Date: 09/08/16 Margaret Briehl

______Date: 09/08/16 Randy Burd

Final approval and acceptance of this dissertation is contingent upon the candidate’s submission of the final copies of the dissertation to the Graduate College.

I hereby certify that I have read this dissertation prepared under my direction and recommend that it be accepted as fulfilling the dissertation requirement.

______Date: 09/08/16 Dissertation Director: Kirsten Limesand 3

STATEMENT BY AUTHOR

This dissertation has been submitted in partial fulfillment of requirements for an advanced degree at the University of Arizona and is deposited in the University Library to be made available to borrowers under rules of the Library. Brief quotations from this dissertation are allowable without special permission, provided that accurate acknowledgment of source is made. Requests for permission for extended quotation from or reproduction of this manuscript in whole or in part may be granted by the Copyright holder.

SIGNED: Alejandro Martinez Chibly

ACKNOWLEDGMENTS

“It takes a village”

There are three questions that every PhD student is continuously faced with throughout the journey of graduate school: Why did you choose this college? Why did you choose that lab? How did you choose your field? As if at this point we had to justify ourselves. Nonetheless, my answer to those 3 questions is the same: Kirsten Limesand. So, I first want to express my most sincere gratitude to Dr. Limesand for her commitment to change lives positively as a mentor, her continuous guidance, motivation, support, and incredible mentorship. “It [Mentorship] is a solemn duty to change lives positively. It is a noble honor to inspire and be there for others. It is an irresistible necessity to have empathy; to understand the situations and the reasons for the actions of others. Real mentoring is less of neither the candid smile nor the amicable friendship that exists between the mentor and the mentee and much more of the impacts. The indelible great footprints the mentor lives on the mind of the mentee in a life changing way. How the mentor changes the mentee from ordinariness to extra-ordinariness; the seed of purposefulness that is planted and nurtured for great fruits; the prayer from afar from the mentor to the mentee; and the great inspirations the mentee takes from the mentor to dare unrelentingly to face the storms regardless of how arduous the errand may be with or without the presence of the mentor” -Ernest Agyemang Yeboah.

Second, I’d like to thank those who paved the way for my success before I was born: my family. Much of my success is only possible because of my mother (Paty) and Father (Agustin). They have supported, motivated, and encouraged me at every step of this journey, not only to achieve success, but to pursue my dreams. I owe them for my life, the many lessons they taught me, and for the many sacrifices made for me and my siblings. I am most grateful for they did not ever let me allow myself to yield to self-doubt. Gracias a mis padres, abuelita, y hermanos, por estar presentes en cada paso que he dado, y por compartir su orgullo y cariño en cada uno de mis andares. Indudablemente, cada uno de mis logros es un reflejo de mis padres, y por tanto es un logro suyo. Comparto con ustedes, mi familia, el fruto de incontables sacrificios. Espero siempre ser su orgullo.

I got additional support from numerous faculty, staff, and students throughout the University, and the Cancer Biology program. Thanks to my committee members for lending me some of their valuable time. Their advice and support is deeply appreciated. I want to specially acknowledge Anne Cione, for playing the role of first-responder, student counselor, conflict mediator, problem solver, and of course, coordinator of the CBIO 5 program. Thanks to Anne, for devoting so much to better our program, and for making our dealings with bureaucracy a lot less of a nightmare.

For making grad school an unforgettable and exciting adventure, I thank all of my friends. In each of them I found the strength behind every step and great moments of joy at the most demanding times of graduate school. To my roommate and gym buddy, Mitzy, who is undoubtedly exhausted of living with my bad jokes and constant teasing, thanks for taking care of me, sister. To Norma, for keeping a promise of friendship in spite of the many miles between us, thank you! Special thanks to my buddy and trainer Eddie for believing in me, pushing my limits, and giving me strength (literally and figuratively). I also want to thank Reyna, Maru, Vero, Karen, Carla, Monx, Karla G, Karla H, Fernanda, Daniela, Janine, Hannah, Claudia, Paloma, Shayan, Felipe, Sergio, Beto, Jon, and Stephen; without their friendship and support I would not be whole.

I also want to acknowledge current and past members of my lab for being the best team I’ve ever been part of. Deborah and Salena, convinced me that getting a PhD was the way to go, and never doubted from the beginning that I would get to the finish line. Maria and Katie, thanks for being much more than lab peers, instead, both are excellent friends and mentors; I’m forever grateful that we were part of this journey together. Zoey, Amy, and Vicky, thanks for the continuous support, for happily and excitedly listening to my stories, and for enabling my coffee addiction. Grace, thanks for being a moral compass in times of need. Also thanks to our newest team player, Maricela, for taking over the business so that I could write my dissertation. Without her help, my mental health was in serious jeopardy. Also, thanks for the champagne! And to my mentees and friends: Asha, Allie, Brenda, Bernard, and Micah, thanks for making me proud!

Finally, the completion of my dissertation was possible thanks to the financial support from CONACyT, the Graduate College at the University of Arizona, the National Institute of Health, as well as the Cancer Biology program. Thank you everyone who made this journey possible.

TABLE OF CONTENTS LIST OF FIGURES ...... 9 ABSTRACT ...... 11 I. LITERATURE REVIEW ...... 13 Clinical Significance ...... 13 Epidemiology...... 13 Treatment of head and neck cancer ...... 14 Salivary Hypofunction and Dry Mouth ...... 16 Radiation-Induced Salivary Hypofunction and Xerostomia ...... 17 a) Direct radiation damage to salivary epithelium ...... 19 b) Radiation effects on associated innervation ...... 20 c) Radiation effects on associated vasculature and stroma ...... 21 Etiology of RT-induced salivary hypofunction ...... 22 DNA damage and acinar cell death ...... 23 Alterations in secretory pathways...... 25 Disruption of salivary gland stem and progenitor cells ...... 27 Clinical management of xerostomia ...... 34 Non-pharmacological approaches ...... 34 Palliative care ...... 34 Therapies under development ...... 37 Protective therapies ...... 37 Regenerative therapies ...... 39 a) Pharmacological approaches ...... 39 b) Gene therapy ...... 41 c) Tissue engineering technology ...... 42 d) Stem cell-based therapy ...... 44 Regulation of stem cell function ...... 46 Extrinsic regulation of stem cell function (the stem cell niche) ...... 47 Intrinsic regulation of stem cell function (Cell polarity) ...... 50 Statement of the problem ...... 54 II. CONCLUSION ...... 55 REFERENCES ...... 60 7

APENDIX I. LABEL RETAINING CELLS IN THE ADULT MURINE SALIVARY GLANDS CONTAIN A DIVERSE POPULATION OF SALIVARY PROGENITORS 95 Introduction ...... 96 Materials and Methods ...... 101 Mice and label retaining assay...... 101 Radiation treatment ...... 101 Immunohistochemistry and Immunofluorescence staining ...... 102 Flow cytometry and cell sorting ...... 104 Real-time PCR ...... 105 Fluorescent in-situ hybridization (FISH) ...... 106 Sphere Assay ...... 106 Secondary sphere Assay ...... 108 Statistical Analysis ...... 108 Results ...... 109 Both acinar and ductal compartments of the salivary glands contain label retaining cells ...... 109 Long-lived LRCs of the salivary glands possess markers of putative progenitor cells ...... 112 Salivary gland LRCs have proliferative potential ...... 116 Salivary gland-derived spheres generate differentiated amylase-secreting cells .... 119 Salivary gland LRCs survive targeted radiation treatment to the head and neck .... 121 Discussion ...... 124 Supplementary material...... 130 APENDIX II. RADIATION-INDUCED COMPENSATORY PROLIFERATION IN PAROTID GLAND LABEL-RETAINING CELLS IS INITIATED BY INHIBITION OF APKC AND NUCLEAR TRANSLOCATION OF YAP...... 132 Introduction ...... 133 Methods ...... 138 Mice and label-retaining assay ...... 138 Radiation treatment ...... 138 Immunohistochemistry and Immunofluorescence staining ...... 139 Saliva collection ...... 141 Data analysis ...... 141 Results ...... 142 8

Radiation induces compensatory proliferation in a unique population of parotid acinar label-retaining cells ...... 142 T560 phosphorylation of aPKC is decreased in salivary gland LRCs following radiation ...... 146 Loss of aPKC induces a hyper-proliferative phenotype in acinar LRCs ...... 148 Acute radiation-induced loss of saliva occurs independently of aPKC ...... 150 Radiation and loss of aPKCpromote nuclear localization of yap1 in acinar LRCs ...... 151 Discussion ...... 155 Supplementary material...... 159 APENDIX III. APKC-DEPENDENT INHIBITION OF NUCLEAR YAP MEDIATES RESTORATION OF SALIVARY FUNCTION WITH IGF-1 ...... 163 Introduction ...... 164 Methods ...... 168 Mice and label-retaining assay ...... 168 Radiation treatment ...... 168 IGF-1 Injections ...... 169 Immunohistochemistry and Immunofluorescence staining ...... 169 Saliva collection ...... 171 Data analysis ...... 171 Results ...... 172 Decreased T560phosphorylation of aPKC in acinar LRCs is associated with chronic loss of function...... 172 IGF-1 regulates nuclear levels of Yap in acinar LRCs in an aPKC-dependent fashion ...... 176 Discussion ...... 180 Supplementary Material ...... 185

LIST OF FIGURES

Figure 2. 1: Label retaining assay in murine salivary glands...... 111

Figure 2. 2: Molecular markers in salivary gland LRCs...... 115

Figure 2. 3: Sphere Assay with murine salivary glands...... 117

Figure 2. 4: Proliferative potential of LRCs...... 118

Figure 2. 5: Differentiation of Salivary gland Spheres...... 120

Figure 2. 6: Effect of radiation on salivary gland LRCs...... 123

Figure 2S. 1 ...... 130

Figure 2S. 2: ...... 131

Figure 3. 1: Acinar LRCs undergo compensatory proliferation 5-7 days post-radiation.

...... 144

Figure 3. 2: Cell populations undergoing compensatory proliferation in acinar compartment of the parotid gland...... 145

Figure 3. 3: T560phosphorylation of aPKC in acinar LRCs decreases after radiation.... 147

Figure 3. 4: Initiation of radiation-induced compensatory proliferation in acinar LRCs is regulated by aPKC-dependent and aPKC-independent mechanisms...... 149

Figure 3. 5: Acute loss of saliva following radiation is aPKC-independent...... 153

Figure 3. 6: Radiation induces nuclear translocation of Yap in acinar LRCs...... 154 10

Figure 3S. 1 ...... 159

Figure 3S. 2 ...... 160

Figure 3S. 3 ...... 161

Figure 3S. 4 ...... 162

Figure 4. 1: Decrease in aPKCT560 in acinar LRCs is associated with chronic loss of function...... 174

Figure 4. 2: Recovery of salivary function with IGF-1 requires aPKC...... 175

Figure 4. 3: IGF-1 reduces levels of nuclear Yap in irradiated acinar LRCs...... 178

Figure 4. 4: IGF-1 inhibition of nuclear Yap is mediated by aPKC...... 179

Figure 4S. 1: IGF-1 does not activate AKT in irradiated acinar LRCs...... 185

ABSTRACT

Radiotherapy is the primary treatment for patients with head and neck cancer, which account for roughly 60,000 annual diagnoses in the U.S. and approximately 500,000 worldwide. About 90% of these individuals receive radiation therapy, and salivary hypofunction and xerostomia occur in 60-85% of these patients due to irreversible damage to the salivary glands.

Current preventative and palliative care fail to improve quality of life, accentuating the need for regenerative therapies. Stem/progenitor-cell based therapies have been proposed to regenerate the irradiated glands; however, the identity of stem and progenitor cells in the adult salivary glands has remained somewhat elusive. Moreover, it is unclear how salivary progenitors respond to radiation and whether they can be stimulated to effectively reinstate salivary function.

The second chapter of the present study describes the development of a label- retaining assay in salivary glands using EdU. The label-retaining cells (LRCs) identified in murine salivary glands have proliferative potential in vitro and expressed markers of putative salivary progenitors, such as Keratin 5, Keratin 14, and c-Kit. Interestingly, LRCs were still present 30 days following radiation, when chronic loss of saliva is evident. The significance of these findings lies in the potential of this model to study the mechanisms that prevent salivary progenitors from maintaining salivary gland homeostasis upon exposure to radiation, which will in turn facilitate the development of regenerative therapies for salivary gland dysfunction. 12

In the following chapter, we show that a unique population of murine salivary gland LRCs undergo compensatory proliferation in response to radiation. The initiation of compensatory proliferation is tightly associated with inactivation of the kinase aPKC and increased nuclear localization of YAP. This part of the study provides novel insights into the regulation of function of salivary gland progenitors, which can be utilized for the development of therapeutic agents to treat salivary hypofunction

Finally, the last chapter describes how the mechanisms found to initiate compensatory proliferation in acinar LRCs as a response to radiation are involved in the regeneration of salivary glands with IGF-1. Administration of IGF-1 post-radiation restores salivary function in mice, but the mechanisms of regeneration are still unknown. Here, we show that IGF-1 requires aPKC to restore saliva production. Further, IGF-1 inhibits nuclear translocation of Yap in an aPKC-dependent fashion. We propose that a tightly regulated balance in the levels of aPKC and Yap in acinar LRCs has to be maintained in order to restore function following radiation.

In conclusion, the findings from this study provide new knowledge in regards to the regulation of function of salivary progenitors during a state of injury (by radiation) and during regeneration (with IGF), and offer potential targets of study for the development of new therapeutics for salivary gland dysfunction. Future studies will determine whether aPKC and Yap can be effectively targeted in salivary progenitors to restore salivary function in head and neck cancer patients who receive radiation therapy.

I. LITERATURE REVIEW Clinical Significance

Epidemiology

In the United States, over 60,000 new cases of head and neck cancer (HNC) are estimated for 2016 (American Cancer Society, 2016). These cancers develop in the oral and nasal cavities, pharynx, larynx, and all of their appendages. The main risk factors associated with HNC are cigarette smoking, tobacco, alcohol, human papillomavirus

(HPV) infection, and Epstein-Barr virus (EBV) infection (Calabrese et al., 2012). HNC prevalence is approximately 2.5 times higher in men than women and account for about

3% of all cancers diagnosed in the United States. Worldwide the number of new HNC cases exceeds 500,000 every year with approximately half of those cases being diagnosed in Europe and about 60,000 in the U.S. (American Cancer Society, 2016).

In addition to localization and stage of the tumor, the high recurrence rates observed in HNC limit the ability to maximize loco-regional control solely with a surgical approach, making radiation therapy (RT) the standard treatment for HNC (Cognetti et al.,

2008; Hinni et al., 2013). A downside of RT is that surrounding non-malignant tissues, such as the salivary glands, are damaged by the radiation leading to the development of long-lasting side effects (Moding et al., 2013; Rathod et al., 2015; Vissink et al., 2003;

Wijers et al., 2002). The main complaint in HNC patients treated with RT is xerostomia, a sensation of severe dry mouth, which is normally associated with loss of saliva or salivary hypofunction (Valdez et al., 1993; Wijers et al., 2002). 14

Loss of saliva and xerostomia are conducive to the development of oral sores, mucositis, difficulty to chew and swallow (dysphagia), speech impairment, and alterations in taste (dysgeusia) (Dirix et al., 2006; Jensen et al., 2003; Kakoei et al.,

2012). In addition, changes in pH and composition of saliva promote the development of dental caries and exacerbate the symptom of xerostomia (Pedersen et al., 2002; Spielman et al., 1981). Ultimately, these complications advance into a life-long problem that promotes a chronic state of malnutrition, further decreasing patients’ quality of life (Jham et al., 2008). The severity of these complications is the main contributing factor to patients’ withdrawal from their cancer treatment.

Treatment of head and neck cancer

Depending on localization and stage of the tumor, treatment of HNC may include surgery in combination with radiotherapy and/or chemotherapy (Collins et al., 2005). Due to the limitations of surgery and to maximize loco-regional control, RT remains the main course of treatment for HNC. Radiation therapy is utilized with both palliative and curative intents, depending on stage of the tumor as well as tolerance to RT toxicity.

Although there are established radiation regimens for HNC, RT doses and fractionations are tailored to each patient to take into consideration the proximity of the tumor to radiosensitive tissues, delineation of optimal treatment margins, and potential toxicity

(Kapanen et al., 2013; Parvathaneni et al., 2012; Robar et al., 2007). Currently available

RT techniques, such as Intensity-modulated radiotherapy (IMRT) and proton therapy

(IMPT), deliver radiation beams more precisely to the tumor while partially sparing 15 surrounding non-malignant tissues to diminish the development of side effects (Braam et al., 2006).

The possibility of surgery is always evaluated as a first-line treatment when it allows for complete removal of the tumor while maximizing preservation of function and minimizing impact on aesthetics and psycho-social consequences (Calabrese et al., 2012;

Cognetti et al., 2008). Failure to fully eliminate the primary tumor during surgery has been considered the main cause of death in HNC patients (Johansen et al., 1976).

Unfortunately, due to the diverse nature and anatomical location of these tumors, it is extremely difficult to establish guidelines or standards in regards to optimal surgical margins that allow for maximal regional control (Hinni et al., 2013; Meier et al., 2005), which limits the ability to eradicate the cancer with surgery alone (Hinni et al., 2013;

Meier et al., 2005). In addition, tumors that involve delicate anatomical areas such as the carotid artery, brachial plexus, and cervical vertebrae, are frequently deemed unresectable

(National Comprehensive Cancer Network, 2016).

Cisplatin-based chemotherapy alongside radiotherapy is the current standard therapy for HNC patients with locally advanced disease (stages III and IVB) (National

Comprehensive Cancer Network, 2016). Alternatively, induction chemotherapy can be used to shrink the tumors before administration of the main chemo-radiation (chemoRT) regime, though it has not been established whether this modality of treatment achieves better survival outcomes over chemoRT alone (National Comprehensive Cancer

Network, 2016). Combination of cisplatin-based chemotherapy with cetuximab, fluorouracil, paclitaxel, docetaxel, or gemcitabine, with or without the addition of RT, is 16 also considered as a first-line therapy for patients with recurrent, unresectable, or metastatic disease (National Comprehensive Cancer Network, 2016).

For the particular case of human papilloma virus (HPV)-positive HNC, due to the considerably younger age of these patients and their superior prognosis, it has been proposed to de-intensify current therapies in order to reduce treatment-related morbidities. For these individuals, replacement of cisplatin with cetuximab, lower doses of radiotherapy, less invasive surgery, and the use of immunotherapy, are being currently evaluated (Kofler et al., 2014).

Salivary Hypofunction and Dry Mouth

Hypofunction of the salivary glands is characterized by partial or total loss

(dysfunction) of the normal functions of saliva, due to a reduction in the amount of saliva produced, as well as alterations in its constituents. Xerostomia is frequently associated with salivary hypofunction and thus it is commonly reported by individuals with loss of saliva. However, the symptom of xerostomia is instigated by several other conditions, such as use of multiple medications, diabetes, dehydration, autoimmune and graft-vs-host disease, and does not necessarily involve salivary hypofunction (Jensen et al., 2010;

Porter et al., 2004). Mild salivary hypofunction can result from infectious diseases, inflammatory processes involving the salivary glands, or the use of certain medications

(Porter et al., 2004). More severe damage to the salivary glands is consequence of therapeutic doses of radiation and Sjogren’s syndrome, in which cases permanent salivary 17 dysfunction is more likely to occur (Jensen et al., 2003, 2010). Individuals with a noticeable deficit in saliva may or may not experience dry mouth (Nederfors, 2000).

Radiation-Induced Salivary Hypofunction and Xerostomia

Due to the high recurrence rates of HNC, achieving maximum loco-regional control with first-line treatment is always desired to improve survival outcomes. This is often accomplished by extending the radiation field beyond the tumor margins at discretion of the medical team, which often results in exposure of surrounding non- malignant tissues, such as the salivary glands1, to doses of radiation that may lie beyond the threshold that allows for preservation of function. It has been estimated that a total dose of 20-30Gy delivered to the salivary glands allows some preservation of function and potential improvement upon completion of treatment (Dreyer et al., 1989). In contrast doses above 50Gy are likely to cause permanent salivary hypofunction (Deasy et al., 2010; Dreyer et al., 1989). This is reflected in approximately 60% of HNC patients receiving conventional RT who complain of chronic xerostomia and suffer some degree of saliva loss (Wijers et al., 2002). In most cases, RT-induced salivary hypofunction fails to recover over time (Kam et al., 2007; Nutting et al., 2011).

The advent of enhanced modalities of RT, such as IMRT and IMPT, made it possible to reduce the dose delivered to salivary glands during the treatment of HNC, which translates to improved salivary function and overall better a patient’s quality of life post-treatment (Bhide et al., 2012; Kam et al., 2007; Nutting et al., 2011; Parvathaneni et

1 With the exception of salivary gland cancer, in which salivary glands are the primary target of treatment. 18 al., 2012; van Rij et al., 2008; Rubin and Casarett, 1968). For instance, in a study comparing IMRT versus conventional radiotherapy for patients with oropharyngeal cancer, it was reported that 55% of individuals treated with IMRT showed persistent reductions in salivary flow 6 months after treatment, compared to 84% in the conventional RT cohort (Braam et al., 2006). In addition, the severity of saliva loss was significantly higher in patients treated with conventional RT than those subjected to

IMRT treatment. The aforementioned improvements in this study could be attributed to the overall lower doses of radiation received by the parotid gland in IMRT-treated patients (33.7 Gy) compared to Conventional RT (48.1 Gy).

Furthermore, based on observations from experiments in rats, a recent study proposed that sparing a critical region of the glands that contains presumed stem cells may be sufficient to preserve salivary function when sparing the whole gland is not feasible (van Luijk et al., 2015). Future clinical studies will be necessary to establish the feasibility of sparing of this critical region in humans and whether it improves quality of life post-therapy.

Despite the promise of IMRT and similar modalities of radiation therapy to maintain some salivary function and potentially improve quality of life, tumor shrinkage as a result of therapy poses a limitation for IMRT, as it may affect the precision of continued treatments (Robar et al., 2007; Rubin and Casarett, 1968). In addition, there is a substantial number of individuals who are not eligible for sparing of the salivary glands, or patients who have already completed their RT regime. These individuals are likely to suffer from chronic loss of saliva, xerostomia, and the gamut of complications associated 19 with these conditions. Therefore, it is imperative to develop restorative therapies to reinstate salivary gland function and achieve measurable improvements in patients’ quality of life.

The severity of xerostomia and the degree of salivary hypofunction correlate with the dose of radiation, the amount of glandular tissue exposed to the radiation field, as well as the extent of damage to salivary epithelium, surrounding nerves, vasculature, and stroma (Dirix et al., 2006; Valdez et al., 1993).

a) Direct radiation damage to salivary epithelium

The salivary epithelium is formed primarily by acinar, ductal, and myoepithelial cells, in addition to the surrounding blood vessels, nerves, and stroma, which are intertwined with the epithelium to form a fully functional gland (Morgan-Bathke et al.,

2014a). The serous and mucous acinar cells are the secretory component of the glands and produce an isotonic amylase-rich watery fluid, and a more viscous mucin-rich saliva, respectively. The ductal system is formed by excretory, striated, and intercalated ductal cells, and its main functions are to modify the electrolyte content of saliva and transport it outwards to the oral cavity (Delporte and Steinfeld, 2006). The combined damage to these structures that results from exposure to RT, leads to a progressive decline in the volume of saliva produced, as well as alterations in the concentration of electrolytes and secretory granules (Niedermeier et al., 1998; Vissink et al., 2003; Wescott et al., 1978).

In patients with radiation-induced sialadenitis, these events are characterized by formation of vacuoles and death of serous acinar cells, followed by damage and loss of 20 mucous acinar cells and development of ductal ectasia (Dreyer et al., 1989). In advanced phases of the response of salivary glands to radiation, the acinar component of the salivary glands continues to deteriorate until most of the secretory cells are lost and ductal metaplasia develops (Dreyer et al., 1989).

Frequently, saliva obtained from HNC patients post-therapy is dense and foamy due to its lower concentration of amylase, higher electrolyte content, and abnormal levels of immunoglobulins A and G, lysozyme, and lactoferrin, which exacerbates the feeling of xerostomia (Cowman et al., 1983; Vissink et al., 2003).

b) Radiation effects on associated innervation

Peripheral nerves provide the necessary stimulus for acinar cells to carry their secretory functions and thus are indispensable for saliva production. Injury to salivary gland innervation could negatively impact the ability of the glands to secrete saliva.

Indeed, observations in submandibular gland biopsies from HNC patients treated with radiation showed ~60% decrease in parasympathetic innervation when compared to non- irradiated controls (Knox et al., 2013).

Animal experiments have confirmed the sensitivity of salivary gland innervation to radiotherapy. For instance, in submandibular glands of rats, a single dose of radiation was shown to be effective in causing edema and loss of secretory synaptic vesicles, which then prevented acinar cell stimulation (Redman, 2008). However, a study suggests that the reduction in parasympathetic nerves is secondary to the damage caused to the salivary epithelium. In this study, radiation administered to embryonic submandibular 21 explants from mice, caused an 80% reduction in the axon number of parasympathetic nerves; however, apoptosis in the salivary epithelium preceded apoptotic events in the parasympathetic ganglion, which led the authors to hypothesize that epithelium-derived neurotrophic factors (i.e. neurturin) are secreted to maintain salivary gland innervation

(Knox et al., 2013).

Even though it has not been tested directly, it is likely that such a dramatic decrease in parasympathetic stimuli contributes to radiation-induced salivary hypofunction and xerostomia.

c) Radiation effects on associated vasculature and stroma

Radiation damage to vasculature activates inflammatory pathways that predispose individuals to vascular disease (Weintraub et al., 2010). The physiological response and extent of damage depends greatly in the type of vessels involved in the injury (Russell et al., 2009). Upon radiation damage, larger vessels suffer from thickening of the intima, proteoglycan deposition and infiltration of inflammatory cells. Small capillaries undergo thrombosis as cells detach from the basal lamina, leading to loss of capillary segments and poor distribution of nutrients (Russell et al., 2009).

Whether injury to the vascular system during treatment of HNC contributes to hypofunction of salivary glands remains a controversial matter. Nevertheless, inflammation within the vascular system as a consequence of radiation is believed to contribute to macrophage infiltration and development of fibrosis in salivary glands

(Redman, 2008; Vissink et al., 2003). Moreover, studies have shown that microvascular 22 density and local blood flow is markedly reduced in salivary glands of mice and minipigs within the first 24 hours after radiation treatment. These studies hypothesized that early and sustained damage to microvasculature negatively impact global nourishment of the salivary glands after RT, compromising its ability to repair damage (Guo et al., 2014; Xu et al., 2010; Zheng et al., 2011).

In addition, damage to the skin, oral mucosa, teeth, periodontal membranes, and bone, may also contribute to worsening of quality of life, but do not necessarily exacerbate salivary hypofunction (Dreyer et al., 1989; Grundmann et al., 2009; Vissink et al., 2003).

Etiology of RT-induced salivary hypofunction

It is well established that most of the biological effects that result from exposure to radiation are due to excessive DNA damage, which occur directly through the breakage of DNA strands by high energy beams, or indirectly through the generation of reactive oxygen species (ROS) by the interaction of ionizing radiation with water molecules (Rubin and Casarett, 1968; Suzuki et al., 2003). These events are more likely to occur at the G2/M transition phase of the cell cycle, at which point cells are more radiosensitive. In contrast, cells are more radio-resistant towards the end of the S phase of the cell cycle when DNA is being duplicated (Rubin and Casarett, 1968; Tamulevicius et al., 2007). Although it is not fully established why, it is possible that the lower concentration of DNA repair enzymes at G2/M phase, and the spatiotemporal distribution and separation of chromosomes during mitosis, contribute to the higher radio-sensitivity 23 observed (Panganiban et al., 2013). Not surprisingly, tissues with high turnover rates are also more radio-sensitive as they are likely to contain a higher percentage of cycling cell populations, whereas slowly dividing tissues are more radio-resistant (Rubin and

Casarett, 1968).

Salivary glands challenge this paradigm as they are exceptionally radio-sensitive in spite of being a highly differentiated tissue with slow cell turnover (Grundmann et al.,

2009; Konings et al., 2005a; De la Cal et al., 2012; Radfar and Sirois, 2003; Rubin and

Casarett, 1968), but the mechanisms of salivary gland radio-sensitivity are poorly understood. Efforts to decipher the etiology of salivary gland sensitivity to radiation have mostly focused on evaluating DNA damage, cellular senescence and apoptosis, disruption of secretory pathways, and inactivation or killing of stem and progenitor cells.

DNA damage and acinar cell death

Radiation can damage the DNA directly by causing breaks in its phosphodiester backbone, or indirectly through the production of ROS that results from interaction of the radiation with water molecules (Azzam et al., 2012; Harfouche and Martin, 2010; Suzuki et al., 2003). By removing an electron from water molecules, water is eventually converted to hydrogen peroxide, superoxide, and hydroxyl radical which quickly react with DNA destabilizing its structure (Azzam et al., 2012).

Several DNA repair mechanisms deal with single strand breaks (SSBs) limiting their functional transcendence. However, when multiple SSBs happen in close proximity, chromosomal breaks can occur leading to double strand breaks (DSBs). These are more 24 difficult to repair and involve mechanisms of low fidelity, that oftentimes lead to aberrant

DNA repair (Panganiban et al., 2013). In response to DNA damage, cells attempt to repair or contain the damage through the activation of cellular pathways that converge in activation of the tumor suppressor p53, which promotes cell cycle arrest, senescence, or apoptosis, to preserve genomic integrity (Moding et al., 2013). If repair fails, the end result is extensive apoptosis, or accumulation of mutations, depending on the extent of damage and the type of cell affected by radiation (Harfouche and Martin, 2010; Suzuki et al., 2003).

In general, preclinical studies have been effective in recapitulating radiation- induced salivary hypofunction, and thus have been extremely valuable in understanding the molecular mechanisms that ultimately lead to chronic glandular dysfunction

(Reviewed in (Grundmann et al., 2009)). Studies in rodents have shown that radiation doses <10Gy induce an acute response characterized by extensive DNA damage that results in activation of p53 and initiation of an apoptotic program, which physiologically translates to loss of up to 60% of salivary function (Avila et al., 2009; Konings et al.,

2005b) and loss of glandular weight. Additional studies using miniature pigs and non- human primates yielded similar results (Li et al., 2005; Stephens et al., 1986).

Importantly, genetic manipulation of apoptosis and cell cycle arrest in rodents by knocking out p53, or constitutively activating AKT, demonstrated that the tumor suppressor p53 is necessary for the salivary glands to elicit an apoptotic response to RT damage (Limesand et al., 2006, 2010). Absence of p53 led to complete inhibition of radiation-induced apoptosis and preservation of function in irradiated mice (Avila et al., 25

2009). In a similar fashion, pharmacological modulation of cell cycle by administration of a cyclin-dependent kinase inhibitor, Roscovitine, prior to radiation, led to transient cell cycle arrest and preservation of salivary function in mice (Martin et al., 2012).

Alterations in secretory pathways

It is well accepted that most of the long-term biological effects provoked by radiation injury are largely a consequence of DNA damage. Nevertheless, ionizing radiation has the potential to interact with and disrupt virtually all cellular components via oxidation (Reviewed in (Azzam et al., 2012)). In particular, cellular membranes are particularly sensitive to lipid peroxidation, which affects their fluidity and permeability

(Azzam et al., 2012). In turn, alterations in cellular membranes may be deleterious for the physiological activity of ion channels, receptors, and enzymes, that depend on their intrinsic interactions with membranes to perform their cellular functions (Ambudkar,

2012).

In salivary glands, it has been reported that radiation causes severe damage to membranous organelles such as the endoplasmic reticulum and the mitochondria, as well as to the cellular membrane (Boraks et al., 2008). Moreover, it has been suggested that radiation has disruptive effects in plasma membrane associations, ion channels and cellular receptors, and calcium signaling, all of which are required for normal cell function and saliva secretion. Normally, cholinergic factors such as acetylcholine from the parasympathetic nerves interact with muscarinic receptors (MR) in salivary glands to promote calcium influx and activation of ion channels, resulting in saliva secretion 26

(Ambudkar, 2012; Knox et al., 2013). In addition, activation of adrenergic receptors is necessary for protein secretion (Reviewed in (Ekström et al., 2011)). Therefore, it is possible that disruption of any of the aforementioned signals could negatively impact saliva production and result in hyposalivation.

Multiple research groups have reported that radiation impairs the ability of the salivary glands to mobilize intracellular calcium through various mechanisms involving the plasma membrane, ion channels, and inflammation (Ambudkar, 2012; Coppes et al.,

2005; Nandula et al., 2013). One such study using rats demonstrated that radiation impaired the ability of parotid glands to mobilize intracellular calcium via disruption of muscarinic receptor-coupled signals through associations of PKCwith the plasma membrane (Coppes et al., 2005). In line with this study, more recent research showed that radiation activated the Transient Potential Melastatin-like 2 (TRPM2) ion channel in cultured human submandibular gland (HSG) cells, and the acinar cells of murine submandibular glands in vivo (Liu et al., 2013). The authors revealed that TRPM2 was activated via generation of ROS and activation of poly-ADP-ribose polymerase (PARP), which led to abnormal Ca2+ influx. Interestingly, knocking out TRPM2 in mice recovered ~60% of salivary function within 30 days post-RT, leading the authors to conclude that radiation-induced activation of TRPM2 may contribute to chronic salivary hypofunction.

Finally, radiation causes deregulation of aquaporin channels (AQPs) in salivary glands of rats, which prevents water movement from the interstitium to the lumen and results in salivary hypofunction (Krane et al., 2001; Takagi et al., 2003; Takakura et al., 27

2007; Vissink et al., 1991). AQPs are water permeable transmembrane channels that regulate water movement towards the luminal side. Consequently, disruption of endothelial AQP1 and acinar AQP5 (which are necessary for saliva secretion) may contribute to chronic hyposalivation (Delporte and Steinfeld, 2006). This is also evident in an AQP5 knockout mouse model, in which pilocarpine-stimulated saliva secretion was decreased by ~60% (Krane et al., 2001). Interestingly, exogenous expression of AQP1 in the ductal cells using gene-therapy approaches, partially restores saliva production and protects the glands from radiation-induced hypofunction (Further described in the sections entitled “Protective therapies” and “Restorative Therapies”) (Baum et al., 2006,

2012; Shan et al., 2005; Teos et al., 2016).

Disruption of salivary gland stem and progenitor cells

Adult stem and progenitor cells (SPCs) have been identified in most adult organs.

Their main function is to continuously renew their homing tissues in response to specific stimuli, such as injury, in order to maintain proper function (Barker et al., 2010; Choi and

Artandi, 2009; Harfouche and Martin, 2010; Insinga et al., 2014). Similar mechanisms of radiation-induced injury to the ones described in the preceding sections, are also damaging to residing SPCs of adult tissues. These mechanisms and the physiological outcomes of radiation damage to SPCs (Summarized in Figure 1.1) are tissue dependent; however, due to the crucial role of SPCs in maintaining tissue homeostasis and promoting repair, disrupting function of SPCs is likely to have long-term functional consequences

(i.e. chronic salivary hypofunction) (Martinez Chibly, 2014; Pringle et al., 2013). 28

As previously described, radiation causes DNA damage, which if left unrepaired, may lead to mutations, senescence, or apoptosis (Azzam et al., 2012). Accumulation of unrepaired DNA damage in SPCs is likely to produce heritable mutations that alter gene expression, which in turn disturb differentiation and self-renewal pathways (Inomata et al., 2009; Insinga et al., 2014; Nicolay et al., 2013; Sokolov et al., 2011). This is often characterized by uncontrolled proliferation, or abnormal states of quiescence (Choi and

Artandi, 2009; Inomata et al., 2009). Consequently, DNA damage to SPCs is one of the mechanisms that leads to carcinogenesis (Harfouche and Martin, 2010; Insinga et al.,

2014). In addition, disrupting normal function of SPCs may have devastating consequences for tissue repair following injury, which has been proposed as one of the mechanisms that lead to permanent radiation-induced loss of saliva (van Luijk et al.,

2015; Martinez Chibly, 2014; Pringle et al., 2013).

SPCs of different tissues respond differently to radiation, with some being able to withstand and repair damage more efficiently than others. For instance, Lgr5+ crypt base columnar (CBC) intestinal stem cells respond like any other cell when are faced with extensive DNA damage: they undergo apoptotic (or mitotic) cell death (Harfouche and

Martin, 2010). However, CBCs repair DNA damage mainly through the high fidelity mechanism of homologous recombination (HR) rather than the low fidelity non- homologous end joining (NHEJ) mechanism of repair, which makes them more efficient at enduring damage and preserving genomic integrity (Hua et al., 2012). Consequently,

CBCs are unusually radioresistant despite having an elevated turnover rate (Hua et al.,

2012; Metcalfe et al., 2014). In line with this, surviving CBCs are capable of naturally 29 regenerating the irradiated intestine (Barker, 2014; Harfouche and Martin, 2010; Hua et al., 2012; Metcalfe et al., 2014). Conversely, stem cells of the skin undergo a process similar to the effects of ageing when exposed to radiation. Upon exposure to radiation,

Keratinocyte stem cells (KSC) are seemingly more resilient to radiation-induced apoptosis, but their colony-forming ability is significantly diminished, and hair cycle is delayed in vivo (Aoki et al., 2013). Along these lines, radiation abrogates the self- renewing potential of Melanocyte stem cells (MSC), which are also present in the skin, forcing them to migrate upwards and differentiate into melanocytes, which is thought to deplete the pool of MSCs (Aoki et al., 2013; Chou et al., 2013; Inomata et al., 2009).

Figure 1. 1: Effects of ionizing radiation on stem and progenitor cells. A) Direct effects of radiation to stem cells: Following radiation, stem and progenitor cells may effectively repair damage by activating DNA repair mechanisms to preserve normal function. Alternatively, extensive damage may lead SPCs to activate apoptotic pathways, which contributes to accelerated ageing by depleting the stem cell pool. Further, unrepaired damage in SPCs that fail to undergo apoptosis is likely to cause premature differentiation or excessive proliferation of SPCs, or accumulation of mutations that promote malignant transformation. B) Indirect effects of radiation to stem and progenitor cells: Radiation injury to the stem cell niche indirectly affects stem cell function. Radiation-induced cell death causes disruption of paracrine signals initiated by the niche and received by the SPCs, which often activate proliferative or apoptotic pathways in SPCs. Cell death activates compensatory proliferation programs, or alternatively may induce apoptosis of SPCs, leading to hyperplasia or accelerated ageing, respectively. Alternatively, surviving SPCs may repair damage and restore tissue function, or become hyperplastic leading to initiation of a tumorigenic process.

This figure was published as part of Palliative Care for Salivary Gland Dysfunction Highlights the Need for Regenerative Therapies: A Review on Radiation and Salivary Gland Stem Cells. J Palliat Care Med. (Martinez Chibly, 2014).

In salivary glands, little is known about the direct effects of radiation on SPCs, mainly as a consequence of the limited knowledge with regard to the identity and function of SPCs in adult salivary glands (Pringle et al., 2013). Pioneering studies aiming to identify salivary SPCs utilized 3H-thymidine or 5-bromodeoxyuridine to label actively cycling populations in the adult salivary glands (Denny and Denny, 1999; Denny et al., 1993;

Man et al., 2001; Takahashi et al., 2001). In these studies, injections of cold thymidine were performed in animals at 10 weeks (or ~2 months) of age, and salivary glands were harvested for analysis at multiple time points ranging from 1 to 42 days following treatment. Based on results from these studies showing elevated proliferation in the excretory ducts, it was speculated that salivary SPCs reside in the ductal system (Konings et al., 2005b; Pringle et al., 2013). In general, salivary SPCs have been identified primarily in submandibular gland based on the expression of Keratin 5, Keratin 14, c-kit, sca-1, and Ascl3, all of which have been demonstrated to have important functions during glandular development (i.e. during morphogenesis) (Knox et al., 2010; Lombaert et al.,

2013, 2008a; Rugel-Stahl et al., 2012; Xiao et al., 2014). However, much less is known about the identity of SPCs in parotid gland, as well as the functional roles they may play during glandular homeostasis and repair in adult salivary glands.

Konings et al. speculated that salivary stem cells were killed or sterilized by radiation treatment, which prevented glandular regeneration (Konings et al., 2005a). In agreement with this, it was recently demonstrated in rats, that by sparing the main excretory duct from the radiation field during RT resulted in improved salivary function

(van Luijk et al., 2015). This region of the glands is believed to contain salivary gland 33 stem cells, which led the authors to hypothesize that inactivation of these stem cells is the cause of radiation-induced chronic salivary hypofunction. In contrast, Keratin 5-positive

(K5+) progenitors of embryonic murine submandibular glands were shown to withstand radiation ex vivo (Knox et al., 2013), albeit glandular regeneration was severely impaired.

Stem cell activity is largely regulated by their interactions with the stem cell niche. These interactions may be in the form of direct adhesions or through the release and subsequent reception of extracellular signals that ultimately regulate cell fate decisions and proliferation (Chen et al., 2013). Thus radiation may also indirectly disturb

SPCs homeostasis by damaging the niche and surrounding cells (Figure 1.1). One such process has been termed “Phoenix rising”, in which apoptotic cell death as a consequence of an external insult, triggers a proliferative response in SPCs to begin wound healing (Li et al., 2010a; Zimmerman et al., 2013). The phoenix rising mechanism seems to involve activation of calcium-independent Phospholipase A2 via Caspases 3 and 7. Downstream reactions lead to the formation of arachidonic acid and prostaglandin E2, which is known to promote proliferation of stem cells (Li et al., 2010a). Besides increased proliferation of stem cells as a result of apoptosis in the niche, this later event may also induce stem cells to undergo cell death (apoptosis-induced apoptosis) via activation of JNK pathway

(Pérez-Garijo et al., 2013). Although it is unclear whether similar mechanisms contribute to radiation-induced salivary gland dysfunction, the study by Knox et al. utilizing irradiated embryonic submandibular gland explants, demonstrated that indeed regeneration was impaired when communication between the parasympathetic nerves

(niche) and Keratin 5 progenitors, was lost (Knox et al., 2013). 34

Clinical management of xerostomia

Non-pharmacological approaches

Non-pharmacological therapies currently under development for the treatment of xerostomia include hyperbaric oxygen (HBO) therapy, acupuncture, electrostimulation, and powered toothbrushes. Of these, HBO is perhaps the more promising alternative as it has shown to improve salivary output in a number of studies, although long-term improvement in xerostomia has been inconsistent (Reviewed in (Fox et al., 2015)).

Acupuncture has yielded mixed results and in general has not met the desired outcomes of salivary output (Simcock et al., 2013; Towler et al., 2013). A recent Cochrane review concluded that while there is low quality evidence that acupuncture slightly increases saliva production, there is insufficient evidence to support that acupuncture can alleviate the symptom of xerostomia (Furness et al., 2013). Similarly, the review concluded that there is insufficient evidence to determine the effects of electrostimulation and powered devices on radiation-induced dry mouth.

Palliative care

Although numerous animal studies have accomplished partial preservation or restoration of salivary function following radiation (described in the following sections), a definitive cure for xerostomia or salivary hypofunction has not been yet developed. To manage the symptom of xerostomia in HNC patients, there are several palliative options available, including saliva substitutes and stimulants, which main goal is to improve quality of life of individuals with salivary hypofunction (Goldstein et al., 2008). 35

In general saliva substitutes and oral lubricants aim to maintain oral health and alleviate the discomfort caused by xerostomia by providing hydration of the oral cavity.

Most commercially available saliva substitutes based on carboxymethyl-cellulose or mucin to increase viscosity. Optimally, a saliva substitute or lubricant should also have remineralizing properties to protect dentins and enamel, and antimicrobial activity to prevent oral infections. In addition to their functional properties, taste, cost, duration of their effects, and presentation (gum, spray, toothpaste, lozenges, gel, mouthwash, etc), are determinant factors for patients’ continual use of these agents. Due to their water- soluble nature, and their continuous loss during swallowing, lubricants are very short- lived and require frequent and repeated re-administration in order to maintain moisture

(Dost and Farah, 2013; Martinez Chibly, 2014; Porter et al., 2004).

Saliva stimulants are particularly useful when there is residual salivary function

(Dost and Farah, 2013; Shiboski et al., 2007). Saliva stimulation can be accomplished locally with chewing gum and lozenges, or indirectly through the use of medication to stimulate the parasympathetic nervous system. Chewing gum supplemented with cariostatic and remineralizing agents have proven to be effective in cases of mild xerostomia, although they have a limited ability to alter saliva composition (Dost and

Farah, 2013). Stimulation of the parasympathetic nervous system through the use of medication, such as pilocarpine, promotes function of exocrine tissues including salivary glands, by promoting the release of acetylcholine and activating cholinergic-muscarinic receptors on salivary acinar cells (Davies and Shorthose, 2007; Fox et al., 1991).

Pilocarpine has been widely used for more severe cases of xerostomia since the early 36

1990s (Fox et al., 1991). However, a recent Cochrane review reported that only about half of the patients from three clinical studies responded to pilocarpine, and that unwanted side effects like sweating, flushing, frequent urination, and headaches, were mild but common. They concluded that there is limited evidence to support the use of pilocarpine to stimulate saliva in patients with radiation-induced xerostomia (Davies and

Shorthose, 2007). Everything considered, saliva substitutes have not consistently demonstrated an improvement of xerostomia in patients with radiation-induced salivary hypofunction (Dost and Farah, 2013).

Therapies under development

Protective therapies

The drug amifostine contains a thiol functional group that acts as a free radical scavenger, which limits the extent of DNA damage caused by RT-generated reactive oxygen species (ROS) (Brizel et al., 2000; Gu et al., 2014). Amifostine is especially attractive as a radioprotectant in the clinic and has been FDA-approved for preventing radiation-induced xerostomia in HNC patients, based on clinical data showing that intravenous administration of amifostine prior to each dose of radiation was able to reduce the incidence of chronic xerostomia, without compromising overall survival

(Brizel et al., 2000). A major caveat of using amifostine is that up to 78% of patients discontinue their treatment due to unwanted side effects like hypotension and vomiting, which are severely exacerbated with concurrent chemotherapy (Rades et al., 2004).

Alternatives to amifostine exist but are still largely in preclinical stages. These include other free radical scavengers, like the stable nitroxide Tempol, and growth factors such as fibroblast growth factor (FGF), keratinocyte growth factor (KGF), and vascular endothelial growth factor (VEGF). Tempol has shown radioprotective effects for salivary glands in mice when administered prior to RT, without affecting tumor sensitivity to radiation (Cotrim et al., 2007a; Vitolo et al., 2004). A study demonstrated that between radiation doses of 12.5-15 Gy, prior administration of Tempol improved salivary outcomes to 70-80% of control values, whereas saliva production in irradiated animals not treated with Tempol was decreased to only 40-50% of unirradiated animals (Vitolo et 38 al., 2004). Similar improvements in saliva production were observed when mice received

Tempol prior to a fractionated RT regime, without affecting tumor sensitivity to radiation

(Cotrim et al., 2007a). Future clinical studies will be necessary to evaluate the effectiveness of Tempol in HNC patients.

On the other hand, growth factors lack the clinical application that other protective and restorative therapies offer for HNC patients with salivary hypofunction due to the lack of selectivity for non-malignant tissues. Systemic distribution of growth factors allows for potential unwanted outcomes that could arise from the interaction with residing SPCs, not only in the tissue of interest, but also in other organs. This is particularly true for younger individuals and cancer patients, in which administration of a growth factor raises concern of relapse or appearance of new malignancies (Chae et al.,

2015; Ogilvy-Stuart and Gleeson, 2004). Nevertheless, gene delivery using adenoviral vectors containing FGF, KGF, or VEGF, has been effective in conferring radioprotection to salivary glands in rodents and minipigs when administered prior to radiation (Cotrim et al., 2007b; Guo et al., 2014; Zheng et al., 2011). Administration of adenoviral vectors expressing either FGF or VEGF via retrograde ductal delivery to the submandibular gland 48 hours prior to radiation, improved salivary flow to ~80-85% of control levels, whereas irradiated mice that did not receive vector, only retained ~35% of saliva production compared to unirradiated mice (Cotrim et al., 2007b). Similar improvements in salivary function were observed in a more recent study using minipigs treated with an adeno-based hybrid vector expressing FGF2, when administered prior to radiation in a fractionated RT regime (Guo et al., 2014). Administration of a comparable hybrid vector 39 expressing KGF also improved salivary function in mice either treated with a single 15

Gy dose of radiation or a fractionated RT regime (6 Gy for 5 days) (Zheng et al., 2011).

Protective therapies are exclusive to newly diagnosed HNC cases, and thus patients currently suffering from radiation-induced salivary hypofunction and xerostomia cannot benefit from them. In addition, the inconsistent efficacy of palliative care in improving quality of life of these individuals underscores the need for restorative therapies (Martinez Chibly, 2014).

Regenerative therapies

Advances in regenerative medicine are largely influenced by our understanding of embryonic development, tissue-specific stem-cell biology, wound healing, and availability of bioengineering tools. Ultimately, the goal of regenerative therapies is to achieve complete functional restoration of a given organ, in order to fight disease, injury, or ageing. In general, recovery of salivary gland function has been approached by means pharmacological treatment, gene-delivery techniques, tissue-engineering technology, and stem cell-based therapies.

a) Pharmacological approaches

Notwithstanding serious obstacles of systemic growth factor administration (Chae et al., 2015; Ogilvy-Stuart and Gleeson, 2004), preclinical studies have evaluated their potential therapeutic use for salivary gland hypofunction. Their utilization may be contingent upon the development of a delivery mechanism that allows targeted 40 administration to the salivary glands (i.e. gene therapy or nanoparticles), still they represent a promising venue for targeting radiation-induced xerostomia.

Results from preclinical studies in mice showed that Insulin-like Growth Factor 1

(IGF-1) and Glial cell line–derived Neurotrophic Factor (GDNF) improve salivary gland function when administered post-radiation (Grundmann et al., 2010; Xiao et al., 2014).

One injection of GDNF (50g/mouse) was given directly into the submandibular gland

(SMG) of mice one day following a single 15 Gy dose of radiation, resulting in improved saliva output, higher sphere forming ability, and increased percentage of functional acini

(Xiao et al., 2014). Similarly, tail-vein injections of IGF-1 administered to mice at days

4-8 post-radiation resulted in elevated salivary flow rates and increased amylase secretion

(Grundmann et al., 2010). The authors of the former study speculated that GDNF acted upon salivary SPCs, based on gene expression analysis showing higher expression of

GDNF in a population of Lin-CD24+c-Kit+Sca1+ cells, which are believed to be salivary

SPCs (Xiao et al., 2014). Although the mechanism of restoration with IGF-1 was not directly evaluated, improved salivary outputs concurred with the resolution of radiation- induced compensatory proliferation, pointing at regulation of proliferation as a potential mechanism of action (Grundmann et al., 2010). Interestingly, treatment with IGF-1 or

GDNF post-radiation did not affect tumor clearance (Victory et al., 2011; Xiao et al.,

2014). In contrast with the aforesaid studies, post-therapeutic injections of KGF in previously irradiated mice had negligible effect in salivary output, and seemed to be more efficient when administered both prior to and following radiation treatment (Lombaert et al., 2008b). 41

Another compound that has been evaluated as a potential therapeutic alternative for radiation-induced salivary hypofunction is the rapalogue CCI-779 (Temsirolimus)

(Morgan-Bathke et al., 2014b). Temsirolimus, which acts through direct and specific inhibition of mTOR, demonstrated to have a similar effect as that of IGF-1 in improving salivary output and amylase secretion while decreasing compensatory proliferation

(Morgan-Bathke et al., 2014b). Perhaps the most attractive feature of temsirolimus is that it is already FDA-approved for renal cell carcinoma, which may accelerate its evaluation in the clinic as a potential therapy for radiation-induced xerostomia.

Lastly, pharmacological activation of the ectodysplasin receptor (EDAR), which is critical for normal development of the salivary glands, showed significant improvements in salivary function of irradiated mice (Hill et al., 2014). Further clinical work will determine if activation of this, or other developmental pathways in salivary glands, are feasible alternatives against salivary hypofunction and xerostomia.

b) Gene therapy

The use of gene delivery technology has been studied in salivary glands by numerous groups with the aim to protect the glands from radiation damage (as explained in the “protective therapies” section), restore salivary function after radiation, treat

Sjögren's syndrome, and as a therapeutic option for gastrointestinal and systemic conditions (Reviewed in (Samuni and Baum, 2011)).

In regards to restoration of salivary function after radiation-induced damage, the most significant progress has been accomplished through the delivery of human 42

Aquaporin-1 (hAQP1) gene with an adenoviral vector (Baum et al., 2010). An initial preclinical study in rats demonstrated that adenoviral delivery of hAQP1 allowed for increased expression of the aquaporin 1 protein in the cell membranes of several cell types in the salivary gland parenchyma. Moreover, salivary flow was 2 to 3-fold higher in irradiated rats that received the hAQP1 gene versus the ones that received the control vector (Delporte et al., 1997). These results, were consistent in a follow-up study using minipigs, which purpose was to test the efficiency of the adenoviral delivery system in a larger animal model (Shan et al., 2005). In the latter study, hAQP1 delivery promoted recovery of salivary flow to about 80% of pre-irradiated values, compared to only 20% in animals receiving a control vector. Finally, a phase I clinical trial in HNC patients who were treated with RT at least 5 years prior to the study, resulted in improved salivary flow (60-540%) in 6 out of 11 individuals who received the AdAQP1 gene therapy

(Baum et al., 2012).

Importantly, although the authors of the study chose an Ad5 vector that does not result in permanent gene transfer, thus limiting issues associated with sustained presence of the virus, there are still experimental caveats and concerns around gene therapy, such as the possibility for gene integration into the host’s DNA, or initiation of an immune response against the vector (Cotrim and Baum, 2008).

c) Tissue engineering technology

One disadvantage of regenerative therapies is that they likely require some amount of intact salivary tissue, which is problematic in patients where extensive fibrosis has developed as a consequence of RT. To circumvent this potential complication, one 43 group proposed to develop an artificial salivary gland (Tran et al., 2006). The authors of the study collected cells from the parotid glands of rhesus monkey and cultured them to evaluate if they could be used to develop a prototype device for in vivo applications.

Their findings show that cells were able to correctly polarize in vitro with correct distribution of ion channels and apical tight junction molecules. Lastly, when cells were transduced with an adeno-based viral vector, they effectively expressed functional water channels for directional fluid movement.

More recently, another research group described a method to artificially generate organ germs derived from the ectodermal placode, with the potential to be transplanted with a regenerative purpose (Nakao et al., 2007). This method was later used to recapitulate the early stages of salivary gland organogenesis in vitro (Ogawa et al., 2013).

A mouse-derived submandibular gland germ was artificially generated and successfully transplanted into a model of salivary gland defect. In this study, single epithelial and mesenchymal cells from embryonic days 13.5-14.5 were isolated and co-cultured to simulate epithelial-mesenchymal interactions that occur during early salivary gland development. The bioengineered submandibular gland formed a fully functional salivary gland able to secrete sufficient saliva in response to pilocarpine and taste stimulation

(Ogawa et al., 2013).

The aforementioned studies provide proof of principle for the use of bioengineering tools to treat salivary hypofunction and xerostomia (Nelson et al., 2013b;

Ogawa and Tsuji, 2015). To achieve clinical applications, however, appropriate cell sources need to be identified. To this end, it has been suggested that the optimal source 44 would be in the form of salivary gland primary or stem cells, which would theoretically be accepted by the host with little or no risk for eliciting an autoimmune response

(Nelson et al., 2013b; Patel and Hoffman, 2014).

d) Stem cell-based therapy

Adult tissues are constantly faced with insults that result from normal biological processes, as well as exposure to exogenous damaging agents. Not surprisingly, adult tissues possess mechanisms of homeostatic regeneration and wound healing that allow them to withstand such continuous stress in order to maintain proper function. This is accomplished through a well-orchestrated process that involves the interaction of signals released from the tissue microenvironment and inflammatory cells, and received by tissue-specific stem and progenitor cells (SPCs) (Arwert et al., 2012; Bergmann and

Steller, 2010; Eming et al., 2014; Fan and Bergmann, 2008; Gurtner et al., 2008; Huh et al., 2004; Karin and Clevers, 2016; King and Newmark, 2012; Lane et al., 2014; Li et al.,

2010a; Tamori and Deng, 2013; Warner and Longmore, 2010; Wells et al., 2006). In turn,

SPCs undergo a ritual of cell division, migration, and differentiation, to compensate for the loss of functional cells (Barker et al., 2010; Simons and Clevers, 2011).

Consequently, adult stem cells are an attractive target for regenerative medicine, and their use has been proposed to promote or accelerate wound healing in a number of ailments including cardiac disease, neurodegeneration, diabetes, and spinal cord injury (Kim et al.,

2013; Liras, 2010; Lunn et al., 2011; Mohsin et al., 2011; Serakinci and Keith, 2006).

In regards to radiation-induced salivary hypofunction and xerostomia, the use of stem cells has been widely proposed as a potential treatment, mainly by means of 45 transplantation, or to use them as a source for generation of a bioengineered organ

(Lombaert et al., 2008a, 2011; Ogawa et al., 2013; Patel and Hoffman, 2014; Pringle et al., 2013; Yoo et al., 2014). For example, Lombaert et al. effectively demonstrated that c- kit+ progenitors from murine submandibular glands had the ability to partially restore salivary function when transplanted into the submandibular glands of irradiated mice

(Lombaert et al., 2008a). In the study, 5 out of 12 animals improved saliva output 60-90 days following transplantation. Xiao et al. recapitulated these experiments utilizing a more defined population of SPCs (Lin-CD24+c-Kit+Sca1+). Injection of 1000 Lin-

CD24+c-Kit+Sca1+ cells into submandibular glands of irradiated mice resulted in significant improvements in saliva production (Xiao et al., 2014).

Undoubtedly, these studies have been instrumental in demonstrating the therapeutic potential of salivary gland SPCs. However, because it is currently unknown whether Lin-CD24+c-Kit+Sca1+ cells (or other salivary SPCs) are damaged by radiation, autologous stem cell transplantation faces similar challenges as bioengineering approaches: a source of salivary SPCs might not be readily available following RT.

Moreover, even if SPCs were to be collected prior to RT to be transplanted upon termination of the cancer treatment, c-kit+ cells lose expression of stem cell markers after three days in culture (Lombaert et al., 2008a), making their culture ex vivo inadequate for any therapeutic application after a regular RT regime for HNC, which lasts approximately 7 weeks. Perhaps a more ideal alternative, is the pharmacological stimulation of endogenous stem cells, which would avoid the need for transplantation. In line with this, it was reported that neurturin-induced stimulation of the parasympathetic 46 nerves, which are necessary for proper function of Keratin 5 (Ker5) progenitors, promoted glandular regeneration in irradiated submandibular explants from mouse embryos (Knox et al., 2013). Although neurturin is not known to have a direct effect on salivary Ker5 progenitors, the study clearly demonstrates that they could be activated through their interaction with the parasympathetic nervous system to favor regeneration.

A setback in achieving this clinically, is the limited knowledge available in regards to the identity of salivary gland SPCs, how their function is regulated, and how they are affected by radiation.

Regulation of stem cell function

As previously mentioned, adult stem and progenitor cells (SPCs) must ensure continuance of proper tissue function by constantly replenishing cells that have fulfilled their life cycle, or have been lost as a consequence of an external insult (Barker et al.,

2010; Hsu and Fuchs, 2012; Yamashita et al., 2010). To achieve this task, SPCs are receptive to signals from their microenvironment (niche) (Choi and Artandi, 2009; Hsu and Fuchs, 2012; Lane et al., 2014; Rezza et al., 2014) and merge these signals with their intracellular machinery to activate a cell division program (Florian and Geiger, 2010;

Martin-Belmonte and Perez-Moreno, 2011; Rodriguez-Boulan and Macara, 2014;

Yamashita et al., 2010). Optimally, adult SPCs generate a clone of themselves and a differentiated cell during each cycle. This process is known as asymmetric cell division.

It allows to effectively restock the functional cells of a given tissue while maintaining a viable pool of SPCs (Barker et al., 2010; Simons and Clevers, 2011). Opposite to this 47 process is the mechanism of symmetric cell division, in which 2 exact replicas of an

SPCs are generated. This process is particularly important during development, when expansion of the SPC pool is required (Morrison and Kimble, 2006). Additionally, symmetric differentiation refers to the generation of 2 daughter cells that are more restricted in their differentiation path than the SPC they derive from. Although asymmetric cell divisions have been described as the main type of proliferation in adult

SPCs, it is likely that all three processes play distinct but necessary roles in the maintenance of homeostasis of adult tissues (Alberts et al., 1997).

Extrinsic regulation of stem cell function (the stem cell niche)

SPCs are not isolated entities, instead, they have intimate connections with their surroundings, from which they obtain the necessary cues to maintain their identity

(Reviewed in (Burness and Sipkins, 2010; Fuchs et al., 2004; Hsu and Fuchs, 2012;

Rezza et al., 2014; Spradling et al., 2001)). The stem cell niche or microenvironment is comprised of cellular and molecular components that provide SPCs with the necessary signals that allow them to thrive, regulating their survival, quiescence, and activation. In general, the components of the stem cell niche are as diverse as the types of SPCs and their homing tissues, yet they are somewhat conserved. The cellular components of most niches are comprised of the SPCs themselves and their progeny, which provide a loop of autocrine and paracrine regulation, and neighboring stromal cells that supply additional paracrine signals. Importantly, these paracrine signals may be received by cells in direct contact with the SPCs or from distant sources within or outside the homing tissue, such as vasculature, nerves, or immune cells. In addition, the extracellular matrix works both as a 48 scaffold and barrier that limits the expansion of SPCs outside of their domain, and regulate their cell fate decisions through direct interactions with integrin molecules

(Rezza et al., 2014).

Communication between SPCs and their niche can be direct through cell-cell contacts, or indirect via secretion of stimulating factors. One good example of the intricate relationship between SPCs and their progeny is the intestine, in which Paneth cells from the niche are in direct contact and offer a necessary stimulus for Lgr5+ stem cells to survive (Clevers, 2013; Clevers et al., 2014). Paneth cells are a differentiated secretory cell type derived from Lgr5+ stem cells, that produces anti-microbial enzymes to protect the intestinal epithelium. In addition to their bactericide functions, direct contact with intestinal stem cells via Notch ligands is required for survival of Lgr5+ cells.

Consequently, genetic deletion of Paneth cells results in concomitant loss of stem cells and disruption of intestinal regeneration (Clevers, 2013).

In addition to survival, SPC quiescence and proliferation are also delicately regulated by niche cells. Hair follicle stem cells (HFSC), for instance, undergo cyclical activation rounds to promote growth of the hair follicles, followed by destruction and rest phases (Alonso, 2006). HFSC reside in a region of the hair follicle known as the bulge, and their function is regulated locally by melanocyte stem cells and keratin-expressing epithelial cells, and distally by the hair germ and the dermal papilla, which are also considered part of the stem cell niche (Rompolas and Greco, 2014). During the growth phase, HFSC proliferation is mediated by activation of the Wnt/-catenin signaling pathway and inhibition of BMP pathway, which occur as a result of paracrine signals 49 produced by the hair germ and the dermal papilla. The latter is also responsible for production of transforming growth factor-β2 (TGFβ2) during the destruction phase of the hair follicle, in which quiescence of HFSC is necessary (Hsu and Fuchs, 2012).

The extracellular matrix (ECM) is in direct contact with many SPCs, functioning as an anchor that prevents expansion outside of their tissues, and regulating cell fate decisions (Watt and Huck, 2013). Similar to the cellular components of the niche, composition of the ECM varies considerably between different tissues, but the overall mechanisms of SPC regulation are somewhat conserved (Gattazzo et al., 2014). SPCs can bind to the ECM via integrins, which trigger intracellular pathways that regulate migration, differentiation, and self-renewal. Additionally, behavior of SPCs during specific events is also influenced by the composition and physical properties of the ECM

(Gattazzo et al., 2014; Watt and Huck, 2013).

Little is known about how the microenvironment modulates stem cell function in salivary glands, perhaps owing to the controversy surrounding the identity of salivary

SPCs in adult glands. Currently, the most objective evidence that an interaction exists between salivary SPCs and their niche, comes from a study in murine submandibular gland explants where it was demonstrated that survival of Keratin 5 progenitors is dependent on the parasympathetic nerves, and removal of the latter component resulted in decreased number of Keratin 5 progenitors, and disturbed morphogenesis of the explant

(Knox et al., 2010). A follow-up study further showed that the parasympathetic nerves were also required for normal function of Keratin 5 progenitors following radiation, and removal of the ganglion disrupted glandular regeneration (Knox et al., 2013). Whereas 50 these results suggest that the parasympathetic nerves are a crucial component of the stem cell niche in the developing submandibular gland, the composition of the niche in adult salivary glands and its interaction with adult salivary SPCs remains to be investigated.

Intrinsic regulation of stem cell function (Cell polarity)

Inside the cell, signals from the environment are translated into specific instructions by the activation of specific signal transduction pathways that govern quiescence and proliferation (Hsu and Fuchs, 2012). Many of these pathways converge in the regulation of cell polarity via the aPKC-Par, Crumbs 3, and Scribble complexes

(Figure 1.2), which in turn dictate whether to expand the pool of stem cells or to differentiate into a more specialized cell (Florian and Geiger, 2010; Martin-Belmonte and

Perez-Moreno, 2011; Yamashita et al., 2010). Perhaps one of the most obvious benefits of a polarized cell is to create a tightly organized intracellular environment, with a differential positioning of organelles along the apical-basal axis, in which cellular processes are optimized (Martin-Belmonte and Perez-Moreno, 2011). One such process is reorganization of cellular membranes and actin filaments during cell division, in which the continuous interaction between adherens junctions, tight junctions, and apical-basal polarity complexes, determine the orientation and symmetry of the mitotic spindle, and the differentiation fate of the daughter cells (Rodriguez-Boulan and Macara, 2014).

The Crumbs 3 polarity complex is formed by the proteins Crumbs 3 (Crb3),

PALS and PATJ, and is necessary for establishment of the apical membrane. The Par- aPKC complex is comprised of atypical protein kinase C (aPKC), partitioning defective 3 51

(PAR3) and 6 (PAR6), and cell division control 42 (CDC42), and is required for appropriate formation of the apical-lateral membrane borders. Finally, the Scribble complex defines the basolateral membrane and is formed by SCRIB, LGL and DLG

(Martin-Belmonte and Perez-Moreno, 2011; Rodriguez-Boulan and Macara, 2014). The many functions of cell polarity complexes in SPCs depend on their correct localization within the cell and properly timed interactions between them. Thus, because of their role in controlling proliferation and differentiation of stem cells, de-regulation of members of the polarity complexes, in particular Scribble and Par-aPKC, have been linked to deficient wound healing, ageing, and tumorigenesis (Martin-Belmonte and Perez-

Moreno, 2011). For instance, overexpression of dominant negative forms of aPKC and overexpression of PAR6, have been associated with hyper-proliferative phenotypes in non-tumorigenic mammary epithelial cells (Nolan et al., 2008; Whyte et al., 2010).

Moreover, loss of polarity is considered a key step in the process of carcinogenesis, and has been associated with increased migration and metastasis (Reviewed in (McCaffrey and Macara, 2011)).

Figure 1. 2: Polarity complexes in SPCs. Normally the Par3-aPKC and Crumbs 3 complexes define the apical membrane while Lgl (Scribble complex) remains at the basolateral domain of the cell. aPKC-Par and Scribble mutually inhibit one another to ensure proper establishment of cell polarity. Crumbs 3 has an extracellular domain and thus and incorporate extracellular signals from the environment into the cell. All 3 complexes interact with each other to maintain proper polarization of the membrane. Moreover, cell polarity complexes are externally regulated by processes of endocytosis and exocytosis, neighboring apoptotic cells, interactions with the niche, and integrin-mediated signaling pathways. Ultimately, activation of aPKC through a sequence of phosphorylations initiates a signaling cascade that regulates stem cell function.

Polarity complexes are central to the asymmetric division process (Goulas et al.,

2012; Martin-Belmonte and Perez-Moreno, 2011; McCaffrey and Macara, 2011;

Rodriguez-Boulan and Macara, 2014; Royer and Lu, 2011), which as previously described, is responsible for maintenance of homeostasis in adult tissues. For instance, a study by Goulas et al. in a Drosophila model showed that the Par-aPKC complex was required and sufficient to drive asymmetric division of adult intestinal stem cells during homeostasis (Goulas et al., 2012). Along these lines, Niessen et al. demonstrated that aPKC regulates self-renewal and differentiation of stem cells in mouse epidermis, and inactivation of aPKC promoted transient proliferation in skin progenitors, depletion of stem cells in the bulge, and altered differentiation of stem cells in the hair follicle and interfollicular epidermis (Niessen et al., 2013). Just as importantly, cell polarity directs stem cell responses during tissue injury events and subsequent wound healing. For example, radiation-induced injury induces excessive proliferation in Drosophila imaginal discs by disrupting the aPKC-Par complex (Warner et al., 2010). Similarly, in the mouse intestine, Lgr5+ stem cells undergo compensatory proliferation in response to radiation injury in order to repair damage (Metcalfe et al., 2014). In this case, loss of aPKC accelerates wound healing by intensifying the proliferative response of the stem cell compartment (Llado et al., 2015).

Statement of the problem

Over 60,000 patients are diagnosed with head and neck cancer each year in the

United States and about half a million worldwide. The main treatment for these patients is radiation therapy, either alone or in combination with chemotherapy and surgery. The majority of individuals undergoing radiotherapy develop chronic salivary dysfunction which fails to improve over time. In turn, the loss of saliva is conducive to a number of complications, such as dysphagia, dental and periodontal decay, chronic oral infections, and malnutrition, all of which contribute to deteriorating quality of life. Currently, there is no cure for the loss of saliva and existing palliative care fails to improve quality of life.

Stem cell-based therapies are an attractive venue for developing effective treatments for radiation-induced loss of saliva; however, the identity of salivary gland stem cells in adult glands remains elusive. More importantly, the effect of radiation on the endogenous stem cell populations of the salivary glands, and whether these cells can be stimulated to restore saliva production is unknown. Therefore, identification of salivary stem and progenitor cells, and understanding how they respond to radiation injury, will facilitate the development of novel therapies for radiation-induced salivary hypofunction.

II. CONCLUSION

Roughly 60,000 individuals are diagnosed with some type of head and neck cancer

(HNC) each year in the United States, resulting in more than 12,000 deaths (American

Cancer Society, 2016). Currently, the standard treatment for HNC is radiation therapy, which maximizes loco-regional control when combined with surgery or chemotherapy

(Cognetti et al., 2008; Hinni et al., 2013).

A big limitation of radiotherapy is that non-malignant tissues, such as the salivary glands, are exposed to radiation during the treatment of the cancer. The damage received by the salivary glands is responsible for the development of long-lasting xerostomia and salivary hypofunction (Moding et al., 2013; Rathod et al., 2015; Vissink et al., 2003;

Wijers et al., 2002), for which there is no cure. As a consequence of loss of saliva, patients often present varying degrees of dysphagia, malnutrition, damaged oral mucosa, tooth and periodontal decay, and persistent oral infections, all of which severely deteriorate quality of life (Dirix et al., 2006; Jensen et al., 2003; Kakoei et al., 2012). The only available treatment alternatives are palliative in the form of saliva substitutes, lubricants, and saliva stimulants. All of them are short-lived and do not consistently improve quality of life (Martinez Chibly, 2014).

Given the ineffectiveness of the current palliative care salivary hypofunction and xerostomia, it is of utmost importance that preventative and regenerative therapies are further developed, in order to preserve or reinstate saliva production and increase quality of life of patients receiving radiotherapy, as well as for those individuals currently living with loss of saliva and xerostomia. 56

Functional regeneration of salivary glands has been achieved in animal studies, but the mechanisms that drive regeneration remain elusive. Furthermore, stem cell-based approaches have been widely proposed for developing regenerative treatments for salivary dysfunction; however, stem cell use is a difficult task due to the limited knowledge in regards to the identity of salivary gland stem and progenitor cells. In the embryonic salivary glands, c-kit+, Keratin5+, and Keratin14+ progenitor cells have been shown to exert important functions for normal development (Knox et al., 2010; Lombaert et al., 2013), but their contribution in the adult glands is mostly unknown.

To circumvent the need to use specific markers to identify salivary gland stem and progenitor cells, we designed a label retaining assay (Chapter I). Label retaining cells

(LRCs) identified with this assay include a heterogeneous population of salivary progenitors, as confirmed by their expression of markers such as Keratin 5, Keratin 14, and c-Kit. Importantly, roughly 70% of LRCs in the acinar compartment did not co- localize with the aforementioned markers, suggesting the possibility that other types of salivary progenitors co-exist in the adult salivary glands. One important finding in the present study is that LRCs are present in the salivary glands at least 30-days post- radiation, when chronic loss of saliva is evident (Grundmann et al., 2010). Moreover, we observed that LRCs do not undergo apoptosis 24-hours post-radiation, when the highest levels of apoptosis are detected in the parotid gland after a 5Gy dose of radiation (Avila et al., 2009; Limesand et al., 2009, 2010; Morgan-Bathke et al., 2014c). These findings argue against the long-standing hypothesis that radiation therapy kills, or sterilizes the 57 residing progenitors of the salivary glands (Konings et al., 2005b), and instead, suggest the potential to stimulate these populations to promote salivary regeneration.

We aimed to further understand how radiation therapy specifically affects salivary gland LRCs, in order to better understand their role in maintaining salivary gland homeostasis after injury. LRCs were found to undergo compensatory proliferation, a process that has been shown to participate in wound healing programs (Bergmann and

Steller, 2010; Blanpain and Fuchs, 2014). Very interestingly, Keratin5+, Keratin14+ and c-Kit+ progenitors in acinar compartment did not significantly contribute to the initiation of radiation-induced compensatory proliferation, and ductal LRCs did not proliferate.

Instead, a yet unidentified subpopulation of acinar LRCs became proliferative at the initiation stages of compensatory proliferation, again suggesting that distinct subpopulations of LRCs might be responsible for maintenance of homeostasis in the adult gland. These observations argue against the current dogma that salivary gland stem cells reside in the ducts, and that these cells are responsible for maintaining homeostasis in the adult salivary glands (Pringle et al., 2013). Rather, our findings show that acinar LRCs

(which include intercalated ductal cells) contribute to replenishing the acinar compartment following radiation without contribution from the major ductal system.

Nevertheless, salivary function is not restored in irradiated salivary glands. It is possible that proliferation of acinar LRCs is not sufficient to restore glandular homeostasis, and that stimulation of additional sub-populations, such as Keratin5+, Keratin14+ and c-Kit+ progenitors is required to achieve functional restoration. This underscores the importance of elucidating which specific cell populations do and do not participate in the radiation- 58 induced compensatory proliferation program and understanding how this process is regulated.

Based on data from other studies that described a similar process of radiation- induced compensatory proliferation which showed that disruption of the Par-aPKC polarity complex promotes proliferation (Llado et al., 2015; Warner and Longmore,

2010; Zimmerman et al., 2013), and particularly, that loss of aPKC accelerates the regenerative process (Llado et al., 2015), we evaluated the significance of aPKC in regulating proliferation of acinar LRCs following radiation. We observed that, similar to its function in mouse intestine, aPKC inhibits proliferation of acinar LRCs in salivary glands through regulation of Yap’s localization. Moreover, decreased levels of aPKCT560 (active form of aPKC) in acinar LRCs and increased nuclear Yap were tightly associated with the initiation of compensatory proliferation (Chapter III), as well as chronic loss of salivary function (Chapter IV).

To further investigate the significance of aPKC and Yap in acinar LRCs to regulate salivary function, we used a previously developed model of salivary regeneration with IGF-1 (Grundmann et al., 2010). Administration of IGF-1 at 4 to 7-days post- radiotherapy leads to partial restoration of salivary function by 30 days following radiation treatment, and full recovery by 60 days (Grundmann et al., 2010). IGF-1 was administered after acute loss of function was evident, and during a stage in which acinar

LRCs initiate a compensatory proliferation program (Chapter III). In our study, recovery of salivary function with IGF-1 is mediated by modulating the levels of nuclear Yes- 59 associated protein (Yap) in acinar LRCs in an aPKC-dependent fashion (Chapter IV).

Moreover, IGF-1 failed to restore saliva in a genetic mouse model with deletion of the

Prkcz gene, demonstrating that aPKC is required for functional restoration of salivary glands with IGF-1.

In summary, we show that a subpopulation of acinar LRCs act as salivary progenitors in the adult gland (Chaper I) and have the potential to replenish the acinar compartment following injury (Chapter III). Radiation therapy induces compensatory proliferation in acinar LRCs by inhibiting phosphorylation of aPKCand promoting nuclear translocation of Yap (Chapter III). Finally, functional restoration of the salivary glands with IGF-1 requires aPKC and involves phosphorylation of the kinase in acinar

LRCs, as well as a reduction in the levels of nuclear Yap (Chapter IV). We propose that a delicate balance in levels of aPKC and nuclear Yap have to be maintained in salivary progenitors in order to restore function of the salivary glands following radiation injury.

This work provides new insights on the mechanisms that prevent salivary regeneration following radiation injury, and offers potential therapeutic targets for restoration of salivary gland function. Future studies will be necessary to determine whether aPKC and Yap, or their downstream effectors, can be targeted directly in salivary progenitors to promote salivary gland regeneration.

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APENDIX I. LABEL RETAINING CELLS IN THE ADULT MURINE SALIVARY GLANDS CONTAIN A DIVERSE POPULATION OF SALIVARY PROGENITORS

Alejandro M. Chibly1, Lauren Querin2, Zoey Harris2, and *Kirsten H. Limesand1,2 1 Cancer Biology Graduate Program, 2 Department of Nutritional Sciences, The University of Arizona. Tucson, AZ 85721, USA

Funding support: NIH R01DE023534 to KHL. AMC was supported by CONACyT, Mexico.

Key words: Progenitors, label retaining cells, LRC, radiation, salivary glands, head and neck cancer, spheres.

* Corresponding Author: Kirsten H. Limesand; 1177 E 4th St. Shantz 421. Tucson, AZ 85721 Phone: (520) 626-4517 Fax: (520) 621-9446 Email: [email protected]

Conflict of interest: the authors declare none

Note: The contents of this chapter were previously published in: Label-Retaining Cells in the Adult Murine Salivary Glands Possess Characteristics of Adult Progenitor Cells. PLoS ONE 9(9): e107893. doi:10.1371/journal.pone.0107893

Introduction

Radiotherapy is the primary treatment for the nearly 500,000 annual cases of head and neck cancer in the world (Castadot et al., 2010; Cmelak, 2012; Lee et al., 2012;

Marcus and Tishler, 2010). Although the goal of radiotherapy is to target the tumor, secondary exposure occurs in surrounding tissues, such as salivary glands and oral mucosa (Grundmann et al., 2009; Redman, 2008). Some of the complications that arise from damage to these normal tissues include acute mucositis, recurrent pneumonia, esophageal dilation, saliva depletion, increased oral infections, and difficulty breathing and swallowing (Cmelak, 2012; Marcus and Tishler, 2010), all of which can last several months or even permanently, contributing to a miserable quality of life.

The FDA-approved drug amifostine has been extensively used in trials as a preventative treatment to ameliorate the side effects that follow radiotherapy (Collins et al., 2005; Dirix et al., 2006; Jensen et al., 2003). Although a reduction of xerostomia has been observed with amifostine, the number of side effects and serious conditions such as hypotension, Stevens-Johnsons syndrome, and hypersensitivity (Gu et al., 2014; Rades et al., 2004), often influence patients’ compliance. Sialagogues are used as palliative care to stimulate saliva flow when partial salivary gland function is retained, but their efficacy highly diminishes with decay of salivary gland function (Jham et al., 2008; Valdez et al.,

1993). Similarly, saliva substitutes are utilized to maintain moisture in the mouth, and to help preserve oral health; however, the use of substitutes is only a replacement therapy and not a cure for xerostomia (Dost and Farah, 2013). Because both preventive and 97 palliative care fail to improve quality of life of patients undergoing radiation therapy, it is necessary to develop regeneration therapies that allow for restoration of salivary gland function.

Adult progenitor cells have been proposed to have significant roles in wound healing responses, tissue homeostasis, and regeneration (Barker et al., 2010; Choi and

Artandi, 2009; George et al., 2011). A previous review has suggested that chronic dysfunction of the salivary glands is due to improper DNA repair in progenitor cells, thereby impairing the ability of salivary glands for self-repair (Konings et al., 2005b). A major problem in the field of adult salivary gland progenitors is that their identity is still somewhat elusive, due to the lack of known specific markers to delineate such populations (Coppes and Stokman, 2011). For this reason, even though salivary gland progenitors have been studied in models of salivary gland development, based on molecular markers identified in other exocrine tissues (Lombaert et al., 2008a; Nanduri et al., 2011; Nelson et al., 2013a), they are limited by the extent of progenitor-specificity of these markers in the adult gland, and the purity of these populations.

Early studies seeking to isolate progenitor cells of developing salivary glands relied on the expression of c-kit, Sca-1, Keratin 5 (K5), Ascl3, and Keratin 14 (K14)

(Bullard et al., 2008; Knox et al., 2010; Lombaert et al., 2008a; Nelson et al., 2013a;

Rugel-Stahl et al., 2012), which have rendered mixed and heterogeneous populations, some of which retain some regenerative potential (Knox et al., 2013; Lombaert et al.,

2013, 2008a) but do not seem to perfectly overlap with one another (Nelson et al., 2013a; 98

Rugel-Stahl et al., 2012), suggesting the existence of multiple progenitor cells in the salivary epithelium.

Studies by Lombaert et al. (Lombaert et al., 2013, 2008a) reported that c-kit+ cells derived from ductal structures of murine submandibular gland, have self-renewal capacity and can differentiate into both acinar and ductal cells in vivo and in vitro. In these studies, c-kit positive cells demonstrated to have the ability to partially restore function of damaged salivary glands after transplantation; however, it remains unclear whether endogenous c-kit+ cells are relevant in wound healing or regeneration of the parotid gland. Additionally, lineage tracing assays have demonstrated that Ascl3 marks progenitor cells in all 3 major salivary glands (Rugel-Stahl et al., 2012), and Keratin 5 is present in submandibular proximal progenitors confined to the basal layer of the ducts

(Knox et al., 2010; Nelson et al., 2013a). Interestingly, Ascl3+ cells were described as a restricted population of progenitors, since they did not generate serous acinar cells and were not precursors to K5+ cells (Rugel-Stahl et al., 2012). Similarly, K5+ and c-kit+ cells share some co-localization during salivary gland development (Nelson et al.,

2013a), but both have been described as different progenitors (Knox et al., 2010;

Lombaert et al., 2013, 2011). Only recently, a study looked at cells that co-express c-kit and K14, which proved important for branching morphogenesis (Lombaert et al., 2013).

However, the role of these populations in homeostasis of the adult parotid and submandibular glands remains to be elucidated.

In order to circumvent the caveats of identifying progenitor cells based on the expression of molecular markers, some techniques have been developed such label 99 retaining assays, which allow for cell sorting techniques and lineage tracing studies

(Duque and Rakic, 2011; Hsu and Fuchs, 2012; Zeng et al., 2010). Label retaining cell

(LRC) assays have contributed in the past to the identification of progenitors in liver (Li et al., 2010b), skin (Terskikh et al., 2012), sweat gland (Leung et al., 2013; Lu et al.,

2012), pancreas (Teng et al., 2007), intestine (Metcalfe et al., 2014; Roth et al., 2012), and other tissues (Maeshima, 2007; Oliver et al., 2004, 2009; Smith, 2005; You et al.,

2011).In the adult salivary glands, LRCs have been found distributed in all parenchymal structures (Kimoto et al., 2007), supporting the idea that multiple progenitors coexist to maintain the complex structure of the adult salivary gland. The significance of these findings lies on the potential to develop new therapies to restore function of the damaged salivary glands, which will be greatly facilitated upon identification of the progenitor cells responsible for maintaining homeostasis and function of the adult salivary glands.

Moreover, it is vital to understand the mechanisms by which radiation therapy corrupts the function of these progenitors so that more efficient targeted therapies can be developed.

In this study we designed a pulse and chase assay utilizing BrdU or EdU injections in a mouse model to detect label retaining cells in the salivary glands. Salivary gland LRCs were present in all parenchymal compartments of the gland, and co-localized with a diverse population of progenitors including c-kit+, K5+, and K14+ cells. Upon culturing in vitro, LRCs initiated sphere formation and differentiated into amylase secreting cells. Finally, we demonstrate that LRCs are maintained following radiation 100 treatment, which makes them extremely attractive for the study and development of regeneration therapies for damaged salivary glands.

101

Materials and Methods

Mice and label retaining assay

All experiments were conducted in FVB mice, both male and female and repeated at least twice and in most cases 3-4 times. For all experiments, date of birth was considered ‘Day 0’ unless otherwise specified. At day 10, mice were given four intraperitoneal BrdU (5-Bromo-2’-deoxyuridine, Roche, Mannheim, Germany) injections at a dose of 3mg per 100g of body weight, or EdU (5-ethynyl-2’-deoxyuridine,

Invitrogen, Carlsbad, CA) injections at a dose of 10mg/100g of body weight 12 hours apart. The chase period for the label retaining assay was 8 weeks, and at this time point the mice were anesthetized via an intraperitoneal injection with Avertin (240mg/kg,

Sigma, St Louis, MO) and euthanized by exsanguination for collection of the salivary glands. All mice were maintained and treated in accordance with protocols approved by the University of Arizona Institutional Animal Care and Use Committee (IACUC). A total of 5 animals (3 males and 2 females) were labeled with BrdU and 46 animals (24 males and 22 females) were labeled with EdU. Mice were distributed to different experiments as indicated in the following sections.

Radiation treatment

Mice were anesthetized via intraperitoneal injection with ketamine/xylazine (50 mg/kg/10 mg/ml) prior to radiation treatment. Radiation treatment consisted of a single dose of 5 Gy targeted to the head and neck region using a 60Cobalt Teletherapy unit from

Atomic Energy of Canada Ltd Theratron-80. The 5 Gy radiation dose was chosen based 102 on our previous work demonstrating the dose caused elevated levels of p53 protein, activation of apoptosis and loss of salivary function (Avila et al., 2009; Limesand et al.,

2009, 2010; Morgan-Bathke et al., 2014c). The remaining body sections of the mice were protected with >6 mm thick lead to avoid systemic effects of radiation. Radiation dosage calculations and maintenance of the cobalt source are conducted by the Experimental

Radiation Shared Service of the Arizona Cancer Center. A total of 10 EdU-labeled mice were treated with radiation, 7 of which (5 females and 2 males) were given the treatment at 4 weeks of age, and the remaining 3 (1 male and 2 females) were treated 24 hours prior to tissue collection at the end of the 8-week chase period. Analysis of %LRCs in irradiated animals was performed using 12 untreated controls (7 males and 5 females) for parotid gland and 6 untreated controls (4 males and 2 females) for submandibular gland, which were compared to specimens obtained from the 7 irradiated animals.

Immunohistochemistry and Immunofluorescence staining

Following dissection, the three major salivary glands were immediately fixed in

10% neutral buffered formalin (Sigma) for 24 hours, transferred to 70% ethanol, and embedded in paraffin. Sections of all major salivary glands were cut to 4μm thickness and processed for standard staining with hematoxylin and eosin by the Tissue Acquisition and Cellular/Molecular Analysis Shared Resource at the University of Arizona. Slides were incubated at 37oC for 20 minutes and rehydrated in Histoclear (National

Diagnostics, Atlanta, GA), graded ethanol (100%-50%) and distilled water. Antigen retrieval was performed placing slides in 1mM citric acid buffer (pH 6.8) and boiling in microwave twice for 5 minutes, plus additional 20 minutes in buffer without further 103 microwaving. For BrdU immunohistochemistry, 0.3% Hydrogen peroxide was used to quench endogenous peroxidases for 15 minutes at room temperature prior to antigen retrieval. Slides were treated as instructed by the manufacturer (Vectastain Elite ABC kit,

PK-6104, Vector Laboratories, Burlingame, CA). Primary Rat monoclonal antibody anti-

BrdU (ab6326, Abcam, Cambridge, England) was used overnight at 4 oC. Positive staining was developed using DAB (Biogenex. San Ramon, CA) for 5 minutes. Slides were counterstained with hematoxylin, dehydrated with graded ethanol washes (50%-

100%) and mounted with Permount (Thermo Fisher Scientific, Pittsburg, PA). For immunofluorescence staining, slides were incubated in 0.02% Triton-x100 solution in 1X

PBS for 15 minutes, followed by three 1X PBS washes of 5 minutes each prior to antigen retrieval. Slides were blocked in 300l of 0.5% NEN and incubated in primary antibody diluted in 1% BSA overnight at 4oC. After 3 consecutive washes with 1x PBS for 5 minutes each, secondary antibody was added for 1 hour at room temperature. Slides were rinsed with 1X PBS and washed with distilled water for 10 minutes. Finally, tissues were counterstained with DAPI (1g/mL) and mounted with a solution of 50% glycerol in 10 mM Tris-HCl (pH 8.0). Fluorescently stained slides were stored at 4 oC for no longer than

5 days until imaging. Images were taken with a Leica DM5500 microscope (Leica

Microsystems, Wetzlar, Germany) and 4 megapixel Pursuit camera (Diagnostic

Instruments, Inc, Sterling Heights, MI) and ImagePro Software. BrdU and EdU-positive cells in parotid and submandibular sections were manually counted from a minimum of 5 fields of view (40X objective) per slide from three slides (three mice) per group. For evaluation of EdU and BrdU efficiency, 5 BrdU-labeled and 5 EdU-labeled mice (both 104 male and female) were analyzed. Cells from acinar and ductal compartments were counted separately and statistical analysis was performed individually as described in the statistical analysis section. Ductal structures were identified based on morphological features (cellular structure enclosing a lumen, example marked with black arrowhead and dashed lines in Figure 1D-E), and were designated as ‘ductal compartment’. The rest of the glandular area was termed ‘acinar compartment’, which includes mostly acinar and myoepithelial cells. Each of these compartments also includes a minority of cells which are not traditional ductal or acinar cells. We used primary antibodies anti Keratin 14

(1:400, PRB-155P, Covance, Princeton, NJ), Keratin 5 (1:400, PRB-160P, Covance),

Monoclonal Anti-Smooth Muscle alpha-Actin (1:500, C6198, sigma), Ki67 (1:200,

12202, Cell signaling Danvers, MA) and Amylase (1:500, clone 1A4, Sigma). Each marker was stained for in tissues obtained from four 10-week old mice (2 males and 2 females), and six 10-day old mice (4 females and 2 males), and each experiment was repeated an average of 3 times.

Flow cytometry and cell sorting

Immediately after dissection, glands from 2 female EdU labeled mice as well as 2 female non-labeled controls were minced in dispersion media containing 1mg/ml of

Collagenase (Sigma), 1mg/ml of Hyaluronidase (Sigma) and 9mM of CaCl2 in Modified

Hanks Balanced Salt Solution (HBSS). Gland preparations were incubated at 37oC for 20 minutes with slight agitation. Cell suspension was pelleted by centrifugation at 100g for 3 minutes and resuspended in 1 mL of 1 M EGTA in HBSS. Cells were put in a rocker for

10 minutes at 37oC to allow for complete digestion. Further, cells were passed through a 105

20 m nylon mesh to separate undigested tissue. Cells were centrifuged at 800 rpm at 4 oC and washed with 1 mL of cold 1xPBS. Cells were pelleted again and resuspended in

500 L of ice-cold PBS. Cells were fixed and stained for EdU following the manufacturer’s instructions (Click-iT® EdU Flow Cytometry Assay Kit, Invitrogen).

EdU+ cells were sorted using the FACSAria (BD Bioscienes, San Jose, CA) and recovered in RNAlater (QIAGEN, Venlo, Limburg) for PCR analysis. Additionally, a

10 L aliquot of the recovered samples was placed in a microscope slide for imaging to confirm presence of EdU+ cells.

Real-time PCR

Gene expression was evaluated from FACS-sorted EdU+ cells. Cells were sorted following FACS procedure described above and collected in RNAlater (QIAGEN) at

4oC. cDNA was obtained directly from cells using FastLane cell cDNA kit (QIAGEN).

Real-time PCR was performed using 5L of cDNA from cells with the appropriate primers. SYBR® Green (QIAGEN) was used for detection. All primers were obtained from Integrated DNA Technologies (Coralville, IA), and their sequences are as follows: amylase (Fwd: 5’-GCTCATCCTTATGGTTTCACACGG-3’, Rev: 5’-

CCAGTCATTGCCACAAGTGCTGTC-3’), aquaporin 3 (aqp3, Fwd: 5’-

CTGCCCGTGACTTTGGACCTC-3’, Rev: 5’-CGAAGACACCGATGGAACC-3’),

Keratin 5 (krt5, Fwd: 5’-GAACAAAGGTGGAGGGAAGA-3’, Rev: 5’-

TGCTGTCCCACCAAATCTTG-3’), Keratin 14 (krt14, Fwd: 5’-

CTGGTGGGCAGTGAGAAAGT-3’, Rev: 5’-CCAGGATCTTGCTCTTCAGG-3’), kit

(Fwd: 5’-TGGTTGTGGTTGTTGTTGTTGTTG-3’, Rev: 5’- 106

GAAGGCTTGTTCCGAAGTGTAGAC-3’), and sox2 (Fwd: 5’-

ATGGACAGCTACGCGCAC-3’, Rev: 5’-CGAGCCGTTCATGTAGGTCTG-3’).

Fluorescent in-situ hybridization (FISH)

Formalin-Fixed Paraffin-Embedded slides from all major salivary glands were obtained from six 10-day old mice (4 females and 2 males) and four 10-week old EdU labeled mice (2 males and 2 females) as described in the previous sections. Slides were pretreated following the manufacturer’s instructions (RNAscope Fluorescent Multiplex,

320850, Advance Cell Diagnostics, Hayward, CA). Slides were incubated with a probe for mouse c-kit mRNA (kit) (NM_021099, Advance Cell Diagnostics, Hayward, CA) for

2 hours at 40oC. Following hybridization, slides were imaged with a Leica DM5500 microscope (Leica Microsystems, Wetzlar, Germany) and 4 megapixel Pursuit camera

(Diagnostic Instruments, Inc, Sterling Heights, MI) using ImagePro Software.

Sphere Assay

A total of 8 EdU-labeled mice (4 males and 4 females) were used for sphere culture across 4 individual experiments. For each experiment, parotid and submandibular glands from 2 mice were collected at week 8 after pulse, and immediately placed in dispersion media containing a mixture of Collagenase and Hyaluronidase (5mg/5mL of modified Hanks solution per gland), 0.1% CaCl2, gentamycin (0.1mg/mL, Life

Technologies, Carlsbad, CA), and fungizone (5 g/mL, Roche). Glands were minced until big clumps dissociated and then incubated at 37oC for 15 minutes with gentle agitation. Cell suspension was then passed through 40 m mesh sterile filters into a new 107 sterile conical tube and centrifuged at 4000 rpm for 10 minutes. Supernatant was discarded and pellet re-suspended in sphere culture media: DMEM/F12 containing streptomycin, penicillin, EGF (20ng/ml, Fisher Scientifics, Waltham, MA), FGF2

(20ng/ml, Sigma), Insulin (10 g/ml, Invitrogen), N2 supplement (1X, Invitrogen),

Dexamethasone (1 M, Sigma) and glutamine (2.5mM). Cells were plated at a density of

400,000 cells per well in ultra-low attachment plates (Corning®, Corning, NY). For collection, plates were visualized under a bright-field microscope to confirm the presence of spheres. Cells were fixed directly in culture prior to collection by adding 1 volume of

10% buffered formalin for 30 minutes at room temperature. Fixing cells at this point helps prevent aggregation of spheres due to further centrifugation steps. After fixation cells were gently centrifuged for 10 minutes to discard the supernatant; cells were then permeabilized with 0.2%TritonX in PBS for 15 minutes at room temperature to initiate the staining procedure. EdU Staining was performed adapting the manufacturer’s instructions (Click-iT® Plus EdU Alexa Fluor® 488 Imaging Kit, Life Technologies,

Grand Island, NY) to stain cells directly in suspension; cells were pelleted by centrifugation at 5000 rpm for 5 minutes and resuspended in EdU click-it cocktail for 30 min at room temperature covered from light. Target-specific staining of spheres was performed in suspension by adding primary antibody diluted 1:200 in 2%BSA for 1 hour at room temperature, followed by incubation in secondary antibody anti-rabbit Alexa

Fluor 594 (A-11037, Invitrogen) at 1:500 dilution in 2%BSA. Between steps, a single wash with PBS was performed. All centrifugation steps after initial collection of spheres were performed at 5000 rpm for 5 minutes. Primary antibodies used were Ki67 (12202, 108

Cell Signaling), and amylase (clone 1A4, Sigma). Amylase-stained spheres were also imaged with the Nikon C1si scanning confocal microscope at the Keck Imaging center at the University of Arizona. Staining of spheres was repeated 4 times for each marker at each collection time point.

Secondary sphere Assay

Primary spheres were grown as described above for 14 days. At this point spheres were mechanically disrupted by passing them through a 28G needle several times and re- plated in fresh media in low-attachment plates. Observations on the day after plating showed that cells were present in small clusters of <5 cells, similar to a previously published report examining cellular clusters that form primary spheres (Lombaert et al.,

2008a). Cells were cultured for an additional 7 days and collected to evaluate the presence of spheres.

Statistical Analysis

An unpaired two-sided T-Test for 2 samples with equal variances was utilized for comparison between BrdU and EdU (n=5 for each group), and to evaluate statistical differences in % of EdU LRCs in irradiated samples (n=7) versus untreated controls

(n=12 for parotid gland and n=6 for submandibular gland). In all cases, data from the acinar compartment were analyzed separately from the ductal compartment. The number of animals utilized for analysis is also indicated under each experiment section as well as every figure legend.

109

Results

Both acinar and ductal compartments of the salivary glands contain label retaining cells

Ten-day old FVB mice were given four EdU (10mg/100g of body weight) or

BrdU injections (3mg/100gr of body weight), 12 hours apart to allow for binding of label to DNA of actively dividing cells (Figure 2.1A). It was determined that 8 weeks was an optimal chase period (Figure 2.1A) as the glands are fully developed at this point, and the number of LRCs in salivary glands represent only a minority of the total tissue, accounting for only about 4.25±0.25% of the parotid gland and 3.07±0.75% of the submandibular gland.

Chromogenic staining of BrdU was preferred initially to count the number of

LRCs present in the salivary glands, since ductal structures were more easily visualized.

However, multiplexing with different antibodies was not always possible with BrdU, and thus EdU was ideal for this application. LRCs were found throughout the gland in both acinar and ductal compartments (Figure 2.1 D,E). BrdU LRCs from 3 mice (2 males and

1 female) were counted manually in both acinar and ductal compartments to calculate the percentage of LRCs based on the total number of cells in each individual compartment.

In parotid gland, 2.34% of the cells in the acinar compartment were LRCs, as well as 6.80% of the cells in the ductal compartment (Figure 2.1 B,D). In submandibular gland, 1.65% of the acinar area were LRCs, while 5.01% of the ductal compartment were

LRCs (Figure 2.1 B,E). Because EdU staining was later used for multiplexing staining, 110 the number of EdU LRCs (Fig 1F-K) from 5 mice (3 males and 2 females) were also counted and compared to BrdU LRCs from 5 mice (3 males and 2 females) to confirm that the labeling efficiency of both compounds was comparable at the specified doses.

The percentage of BrdU LRCs was not statistically different from the percentage of EdU

LRCs (n=5, p=0.81 for acinar LRCs and n=5, p=0.25 for ductal LRCs) (Figure 2.1C), which validates the use of both compounds interchangeably in our study.

111

Figure 2. 1: Label retaining assay in murine salivary glands. A) Label Retaining Assay. At 10 days after birth (P10), FVB mice were pulsed with 4 doses of EdU (100 mg/kg) or BrdU (30 mg/kg) 12 hours apart. Tissue was collected from 10-week old animals. B) LRCs from 3 mice were manually quantified per salivary gland compartment (acinar and ductal). Data are expressed as percentage from the total of cells in each individual compartment for both parotid and submandibular glands. C) Comparison of labeling efficiency between EdU and BrdU. Data are shown as percentage of LRCs per individual compartment. A 2-sided unpaired T-test was performed for analysis (n = 5 per group). D–E) Representative images of parotid and submandibular glands of BrdU-pulsed animals. LRCs in the acinar compartment (a) are shown with white arrowheads. LRCs in ductal compartment (d) are pointed with black arrows. Example of ductal compartment is delineated by dashed line and pointed with black arrowhead F–K) Representative fluorescent images of salivary glands from EdU-pulsed animals. EdU LRCs are shown in green and DAPI in blue.

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Long-lived LRCs of the salivary glands possess markers of putative progenitor cells

Progenitor cells have long believed to be localized to the ductal structures of the salivary glands (Denny and Denny, 1999; Denny et al., 1993), but we found LRCs distributed in all parenchymal structures. As an initial screening test to determine whether salivary gland LRCs had the potential to be progenitor cells, EdU+ cells from EdU- labeled FVB mice were sorted at the end of the chase period by flow cytometry (Figure

2S.1 A-K). Next we measured the expression of putative stemness-related genes in the salivary glands by real time PCR, such as sox2, kit, and aqp3, as well as the acinar differentiation marker amylase. Edu+ cells from parotid glands were enriched in markers sox2, kit and aqp3, and had lower amylase expression in comparison to EdU- cells

(Figure 2S.1 L).

In addition to c-kit, Keratin 5 (K5) and Keratin 14 (K14) have also been associated with salivary gland progenitors during development (Knox et al., 2010, 2013;

Lombaert et al., 2013); therefore, we aimed to determine whether salivary gland LRCs expressed these markers in vivo. Immunofluorescence staining of tissue sections from 10- day old FVB mice was performed to confirm the presence of K5 and K14 in parotid and submandibular glands (Figure 2.2, 2S.2). Results were compared to immunofluorescence staining of adult tissue sections (10-week old mice) of EdU labeled animals.

In parotid glands of 10-day old mice, K14 was found in both acinar and ductal compartments (Figure 2.2 A, 2S.2 A), but it was almost exclusively found in ductal 113 structures in 10-week old mice, where it co-localized with ductal LRCs (Figure 2.2B, yellow arrowhead, 2S.2B). A small group of LRCs in the acinar compartment co- localized with weakly stained K14+ cells (Figure 2.2B, white arrowhead, 2S.2B). In submandibular gland, K14 was found mostly in the excretory ducts of both 10-day old and 10-week old animals, with weak staining of cells in the acinar compartment (Figure

2.2C-D, 2S.2C-D). A small subset of submandibular LRCs co-localized with K14+ cells in the acinar compartment (not shown), but virtually no co-localization with ductal K14+ cells was observed (Figure 2.2D, 2S.2D). K5 was found in the basal layer of ducts, as well as distributed throughout the acinar compartment in 10-day old mice (Figure 2.2E, I,

2S.2E, I). In adults, basal localization of K5 was conserved (Figure 2.2J, 2S.2J), and K5+ cells in the acinar compartment had myoepithelial-like morphology (Figure 2.2F, 2S.2F).

K5+ cells often co-localized (20.13% ± 2.47%, n=5) with LRCs in the acinar compartment of parotid and submandibular glands (Figure 2.2F, H, white arrowhead,

2S.2F, H), while ~1% of the ductal K5+ cells showed co-localization with ductal LRCs

(Figure 2.2J, L, yellow arrowhead, 2S.2J, L). Staining with anti-Smooth Muscle alpha

Actin (SMA) showed the presence of myoepithelial cells in both glands at 10 days of age

(Figure 2.2M, O, 2S.2M, O), and demonstrated the myoepithelial nature of a number of

LRCs in the acinar compartment of both glands in 10-week old animals (Figure 2.2N, P,

2S.2N, P). Fluorescence In-Situ Hybridization (FISH) was used for detection of kit mRNA in tissue sections. In parotid gland, kit RNA was found distributed throughout the gland in 10-day old animals (Figure 2.2Q, 2S.2Q), while it was expressed mostly in small ducts of the adult gland (Figure 2.2R, 2S.2R). In submandibular gland kit was mainly 114 found in small ducts of both developing and adult glands (Figure 2.2S-T, 2S.2S-T) and weakly distributed to the rest of the tissue. LRCs often showed kit RNA expression in small ducts and acinar compartment cells of both parotid and submandibular glands

(Figure 2.2R, T, 2S.2R, T), while no co-localization was observed in major ductal structures. These results confirmed the observations from the initial screening, which indicated that EdU+ from adult mice cells were enriched with c-kit (Figure 2S.1L).

Combined, these findings suggest that salivary gland LRCs are long-lived cells that are present in the salivary glands at postnatal day 10, and conserve their molecular signatures, such as the expression of K5, K14 and c-kit, as well as their localization.

Additionally, co-localization of the LRCs with K5, K14, SMA and c-kit in multiple gland structures strongly suggests that LRCs are a heterogeneous population of putative progenitor cells.

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Figure 2. 2: Molecular markers in salivary gland LRCs. Representative images of parotid and submandibular glands of 10-day old and 10-week old animals stained for Keratin 14 (A–D), Keratin 5 (E–L), Smooth Muscle alpha Actin (M–P). Q–T) Fluorescence in Situ Hybridization for kit mRNA. EdU LRCs are shown in green, DAPI in blue, and all other markers in red. White arrowheads point at co-localization of each marker with the LRCs in the acinar compartment. Yellow arrowheads point at co-localization of each marker with the LRCs in the ductal compartment. Full size images of every panel are shown in Figure 2S.2. 116

Salivary gland LRCs have proliferative potential

Although co-localization of LRCs with c-kit, K14 and K5 was evident, it is unknown whether these markers are specific for progenitor cells in the salivary glands, and despite their role during development, their contribution to the pool of adult progenitor cells is unclear. Moreover, although label retaining cells are long-lived cells that endure long periods of time in a state of quiescence, they are not necessarily progenitor cells (Duque and Rakic, 2011; Hsu and Fuchs, 2012). The sphere culture assay has been largely used as a tool to identify stem cells based on their capacity to self-renew and differentiate in vitro (Pastrana et al., 2011). Therefore, to confirm that LRCs are progenitor cells, we sought to determine the capacity of salivary gland LRCs to form spheres in vitro under low-attachment conditions. Since LRCs co-localized with putative progenitor markers in vivo, we hypothesized that LRCs have the potential to expand and differentiate in vitro to form spheres.

Spheres from parotid and submandibular glands were cultured as explained in the methods section. Spheres were first detected at day 2 after culture (Figure 2.3 A,B Day2) and reached their maximum size around day 5 (Figure 2.3). At day 5, lots of cells were detaching from the spheres (Figure 2.3A,B Day5) and cell death was greatly increased

(data not shown). Strikingly, essentially every sphere contained at least one EdU LRC

(Figure 2.4A,D), and secondary spheres grown from dissociated primary spheres also contained EdU+ cells (Figure 2.4E-F), suggesting that LRCs are involved in sphere formation. 117

To confirm LRCs have proliferative potential, we looked for co-localization of

LRCs with the marker of proliferation Ki67 throughout sphere formation. Co-localization of EdU+ cells with Ki67 was observed during early stages of sphere formation (Figure

2.4 A-C), confirming that LRCs have the potential to proliferate in vitro. At later time points in culture, non-LRCs, are highly proliferative, while LRCs seem to return to a state of quiescence (Figure 2.4D).

Figure 2. 3: Sphere Assay with murine salivary glands. Representative microscope images of spheres grown from submandibular (A) and Parotid (B) glands from 10-week old mice

118

Figure 2. 4: Proliferative potential of LRCs. A–C) Dual staining for EdU and Ki67 in parotid-derived spheres from 10-week old mice. Co-localization is shown with white arrowheads. D–F) Secondary spheres stained for EdU. G–I) Dual staining for EdU and Ki67 in parotid gland tissue sections of 10- week old mice. Zoomed region in yellow square is shown in I’. In both panels, Ki67 is shown in red, EdU in green, and DAPI in blue.

119

Salivary gland-derived spheres generate differentiated amylase-secreting cells

Because LRCs showed multiple features of progenitor cells, including the ability to expand and form spheres in vitro, we hypothesized that LRCs had the capacity to generate differentiated salivary gland cells. To evaluate the differentiation capacity of

LRCs, we performed dual immunofluorescent staining of EdU and amylase on spheres derived from parotid gland from EdU-labeled mice. Since the beginning of sphere formation, amylase was detectable in only a few cells (Figure 2.5A), but with longer periods in culture, there was an increase in the number of Amylase+ cells within the spheres (Figure 2.5B-D). Importantly, LRCs did not co-localize with Amylase+ cells in vivo (Figure 2.5G’) or in vitro (Figure 2.5C-D), and only traces of EdU were detected in some Amylase+ cells within the spheres (Figure 2.5C’, white arrow). This may represent some EdU+ cells that have proliferated early during culture (and therefore diluted the

EdU label) and subsequently have undergone differentiation at later stages of culture. The loss of acinar cells is partly responsible for salivary gland dysfunction; therefore, the capacity to generate differentiated amylase-secreting cells has therapeutic potential.

120

Figure 2. 5: Differentiation of Salivary gland Spheres. A–B) Amylase staining (red) of parotid-derived spheres at days 2–3 in culture. C–D) Confocal images at Z = 0.5 um and Z = 6 um of double staining for amylase (red) and EdU (green) at day 4. Areas in yellow dashed squares are shown in C’ and D’. White arrow points at an amylase-positive cell with traces of EdU. Glands were obtained from mice at 10 weeks of age. E–G) Double immunofluorescence staining for Amylase (red) and EdU (Green) of parotid gland of 10-week old mice. White arrowhead points at LRCs in the acinar compartment; yellow arrowhead points at LRCs in ductal structures.

121

Salivary gland LRCs survive targeted radiation treatment to the head and neck

As mentioned before, it has been postulated that radiation treatment causes loss and/or sterilization of salivary progenitor cells, impairing the ability of the tissue for self- repair (Konings et al., 2005b). Because LRCs comprised a mixed population of cells with progenitor capacity, we decided to evaluate the effect of targeted radiation treatment upon these populations. For this purpose, a group of EdU-labeled FVB mice, was subjected to a single 5Gy dose of targeted radiation of the head and neck region at week 4

(Figure 2.6A). A control group was labeled with EdU but was excluded from the radiation treatment for comparison. Mice were euthanized at week 10 and salivary glands were collected for analysis by immunofluorescence.

EdU staining of parotid and submandibular glands (Figure 2.6B-C) revealed no differences in the percentage of LRCs present in either acinar or ductal compartments in irradiated samples when compared to untreated controls (Figure 2.6D-E) (p>0.05 in all cases by unpaired 2-sided T-Test for 2 samples with equal variances, see radiation section of methods for number of samples). This particular finding indicated that the number of

EdU+ progenitor cells was maintained long after radiation treatment. In addition, dual immunofluorescence staining of EdU and Cleaved Caspase 3 was performed on tissues collected 24 hours-post radiation treatment to evaluate apoptosis of LRCs at this particular time point. No co-localization between activated Caspase 3 and EdU was observed in either parotid or submandibular glands of 3 different mice (Figure 2.6G-J). 122

The preservation of LRCs following treatment, combined with their maintenance of proliferative potential in vitro, as well as the generation of amylase-secreting acinar cells within spheres, makes LRCs a valuable target for development of regenerative therapies, which could be applied to head and neck cancer patients undergoing salivary gland dysfunction as a consequence of radiotherapy.

123

Figure 2. 6: Effect of radiation on salivary gland LRCs. A) Experimental setup. A single 5 Gy dose of radiation was given at week 4 to EdU-pulsed animals (n = 7). Tissue was collected at week 10. Representative images of EdU staining of parotid (B) and submandibular (C) glands are shown for irradiated animals and untreated controls. EdU positive cells were quantified manually per individual compartment for both treatments and expressed as percentage of LRCs per compartment for both glands (D–E). P values were obtained with 2-sided unpaired t-test per compartment (n = 7 for irradiated groups, n = 12 for parotid untreated control group and n = 6 for submandibular untreated control group). F) Experimental setup. 5 Gy dose of radiation treatment was given 24-hours prior tissue collection to EdU-pulsed animals (n = 3). G–J) Immunofluorescence staining of Activated Caspase-3 (red) and EdU (green) in parotid and submandibular glands. No co-localization is observed.

124

Discussion

Radiation-induced salivary gland dysfunction is the cause of miserable quality of life in the roughly 50,000 annual cases of head and neck cancer in the U.S (Cmelak,

2012; Marcus and Tishler, 2010). Here we present a model of label retaining cells in murine salivary glands, which can be used in the study of regenerative therapies for irradiated salivary glands. We demonstrate that label retaining cells (LRCs) encompass multiple populations of salivary progenitors, which have the ability to differentiate into amylase-secreting cells. Most importantly, we are the first group to demonstrate that the number of salivary gland progenitor cells is maintained long term following radiation, which makes them a valuable candidate for restorative therapies.

We were able to design a label retaining assay for the salivary glands in which we obtained comparable numbers of LRCs to those previously shown by Kimoto et al.

(Kimoto et al., 2007). Previous studies have used BrdU as a means to identify label retaining cells in several tissues, salivary glands included (Adams and Oxburgh, 2009;

Chen et al., 2014; Kimoto et al., 2007; Li et al., 2010b; You et al., 2011). Currently, more flexible alternatives are available, such as EdU incorporation, which suppresses the use of antibodies and thus eliminates the problem of cross-reactivity with multiplexing staining

(Lu et al., 2012; Zeng et al., 2010). We chose postnatal day 10 as the pulse (labeling) time point based on a previous study of label retaining cells in the submandibular gland

(Kimoto et al., 2007). The rationale was that at P10, progenitor cells are still actively proliferating, allowing for incorporation of EdU; if cellular markers expressed at P10 are 125 then found in co-localization with LRCs in adult salivary glands, it suggests that shortly after P10, these cells became and remained quiescent throughout the 8-week chase period, consistent with progenitor cell behavior.

We demonstrate here that LRCs are salivary gland progenitors, based on their co- localization with molecular markers associated with salivary progenitors and their maintenance of proliferative potential in vitro. Co-localization with c-kit was expected, since earlier studies demonstrated that c-kit+ cells were important progenitors during development (Lombaert et al., 2013, 2008a) and had partial regenerative potential upon transplantation to irradiated salivary glands (Lombaert et al., 2008a). However, it was surprising that co-localization between LRCs and c-kit was not observed in major ductal structures, where other progenitor markers such as K5 have been reported (Knox et al.,

2010; Nelson et al., 2013a). This could likely signify that LRCs encompass a heterogeneous population of progenitor cells, distinctly localized to different compartments of the salivary glands. Indeed, we found Keratins 5 and 14 to be expressed in the major ducts of parotid and submandibular glands, but they rarely co-localized with

LRCs (~1%). Surprisingly, about 20% of the K5+ cells in the acinar compartment were

LRCs. K5+ cells have been described as proximal progenitors during submandibular gland development (Knox et al., 2010), but in our study, the abundance of K5+ LRCs in the acinar compartment suggests that in the adult glands, a subset of K5+ cells might represent a population of progenitors restricted to a different lineage, possibly acinar or myoepithelial. K14 on the other hand, was highly expressed in ductal cells of the parotid gland, where kit mRNA was also expressed. Importantly, K14 has demonstrated a very 126 important role in salivary gland organogenesis in combination with c-kit (Lombaert et al.,

2013), and its co-localization with LRCs in a compartment where kit RNA is highly expressed, suggests that K14+ cells in adult parotid gland might also be important for gland homeostasis. Interestingly, similar to K5, K14 was found in the basal layer of major ducts, but didn’t co-localize with LRCs in that compartment. K14+ LRCs cells were found in small ducts of parotid, and sparsely localized in submandibular gland.

These patterns of LRCs co-localization with the aforementioned markers support the existence of multiple progenitors within the salivary epithelium. In concordance with this, a study of murine sweat glands reported that LRCs comprised three populations of adult progenitors, all of which contribute to homeostasis of the glands (Lu et al., 2012).

In addition to the presence of progenitor markers in LRCs, their contribution to sphere formation further supports their role as adult progenitor cells. The sphere assay is based on the ability of stem/progenitor cells to proliferate, self-renew and differentiate in suspension (Gil-Perotín et al., 2013; Pastrana et al., 2011). The sole presence of LRCs within the spheres at every time point, but most significantly, in secondary spheres, indicates that these are cells with high capacity to survive the in vitro environment in spite of lacking an extracellular matrix. Sphere assays, have been employed in a number of tissues, including pancreas (Rovira et al., 2010), mammary glands (Dontu et al., 2004), and neural tissue (Bao et al., 2006; Gil-Perotín et al., 2013), in which spheres are enriched with cells that harbor self-renewal and differentiation capacity.

Moreover, while salivary glands are known to hardly proliferate in vivo

(Grundmann et al., 2010), LRCs became actively proliferating cells upon culture in 127 sphere-forming conditions. A similar behavior has been observed in other models of label retaining cells in a variety of tissues (Metcalfe et al., 2014; Oliver et al., 2004, 2009; You et al., 2011). Importantly, proliferation of LRCs in these tissues was often related to damage repair and homeostasis. Oliver, J. et al. (Oliver et al., 2004, 2009) demonstrated that LRCs in the renal papilla were directly involved in renal repair upon a transient ischemic event. Similar to our study, LRCs in the papilla were essentially quiescent in homeostasis but showed extensive proliferative potential otherwise. In their study, proliferation was triggered by tissue injury, whereas in our model it was activated upon culture in vitro. One more study in lacrimal glands (You et al., 2011), also showed proliferation of lacrimal LRCs, which was initiated by induction of a severe inflammatory response. It is also noteworthy that salivary gland LRCs did not show signs of amylase expression both in vivo and in vitro, suggesting an undifferentiated phenotype.

The most important finding in the present study is that the percentage of salivary gland LRCs in the submandibular and parotid salivary glands is maintained after radiation. It has been previously demonstrated that levels of apoptosis in the parotid gland peak at 24 hours following radiation (Avila et al., 2009; Limesand et al., 2009, 2010;

Morgan-Bathke et al., 2014c); therefore the level of apoptotic LRCs was evaluated at this time point. We observed that LRCs do not undergo apoptosis at this particular time after radiation in both parotid and submandibular glands. However, due to rapid clearance of apoptotic cells in vivo, the possibility remains that some LRCs could undergo apoptosis prior to the analyzed time point. It has been speculated that radiation therapy kills, or sterilizes the residing progenitors of the salivary glands (Konings et al., 2005b), but this 128 was never addressed experimentally. Transplantation assays have shown that c-kit+ progenitors have potential to partially restore salivary gland function; however, it is unclear whether the endogenous c-kit+ cells of the irradiated gland respond in any way to radiation treatment. In contrast, a more recent study (Knox et al., 2013) reported that K5+ progenitors survived in explants of murine embryonic SMG after ex vivo radiation treatment and retained the ability to regenerate the gland upon administration of

Neurturin. The same study found that K5+ cells remained after radiation in human biopsies from adult salivary glands, and concluded that maintenance of parasympathetic innervations after radiation could aid in regeneration of glandular function. An explanation for maintenance of K5+ progenitors after radiation is that they are in fact resistant to radiation and escape apoptosis, or that the loss of endogenous K5+ cells triggers a proliferative response in the surviving progenitors, maintaining a constant number. Whether these observations hold true for progenitors of the parotid gland remains to be elucidated. Importantly, however, LRCs are maintained in both submandibular and parotid salivary glands after radiation (Fig 6D-E), and similar to K5+ progenitors, there could be potential for regeneration upon administration of the right stimuli.

In contrast with our study, it was very recently reported that Lgr5+ LRC stem cells underwent apoptosis due to extensive radiation-induced DNA damage, but the surviving LRCs were able to restore function (Metcalfe et al., 2014). This emphasizes the importance of understanding the mechanisms of radiation damage to tissue-specific stem cells. Thus, it is relevant to further study how radiation therapy specifically affects 129 salivary gland LRCs, to facilitate the development of regenerative therapies for patients undergoing radiation-induced xerostomia.

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Supplementary material

Figure 2S. 1 A–C) FACS analysis of 2 Edu-labeled mice (B–C) and an unlabeled control (A). P2 is the population of EdU+ sorted cells. D–E) Microscope images from non-sorted cells from unlabeled control. F–K) Microscope images of sorted EdU+ cells from labeled mice. L) RNA analysis comparing gene expression of EdU+ (P2) sorted cells versus EdU− (P3) sorted cells.

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Figure 2S. 2: Full size representative images of immunofluorescence staining for Keratin 14 (A–D), Keratin 5 (E–L), and Smooth Muscle alpha Actin (M–P) in both glands from 10-day old and 10-week old animals. Q–T) Images of FISH for kit mRNA in both glands from 10-day old and 10-week old animals. Yellow squares in all images indicate the corresponding zoomed-in areas shown in Figure 2.2.

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APENDIX II. RADIATION-INDUCED COMPENSATORY PROLIFERATION IN PAROTID GLAND LABEL-RETAINING CELLS IS INITIATED BY INHIBITION OF APKC AND NUCLEAR TRANSLOCATION OF YAP.

Alejandro M. Chibly1, Sourav Ghosh2, and *Kirsten H. Limesand1,3 1 The University of Arizona, Cancer Biology Graduate Program; Tucson, AZ 85721, USA. 2 Yale University, Department of Neurology, New Haven, CT 06511, USA 3Department of Nutritional Sciences, The University of Arizona Tucson, AZ 85721, USA

Funding support: NIH R01DE023534 to KHL. AMC was supported by CONACyT, Mexico.

Key words: Salivary glands, label retaining cells, LRC, radiation, progenitors, head and neck cancer, aPKC, Yap1.

* Corresponding Author: Kirsten H. Limesand; 1177 E 4th St. Shantz 421. Tucson, AZ 85721 Phone: (520) 626-4517 Fax: (520) 621-9446 Email: [email protected]

Conflict of interest: the authors declare none

Note: The contents of this chapter were previously submitted for publication to Cell Death and Disease.

133

Introduction

Head and neck cancers, are diagnosed in approximately 60,000 individuals every year in the United States (American Cancer Society, 2016), and about 90% of these cases receive radiation therapy. Nearly all of these patients suffer from adverse side effects of radiation during the course of treatment, and 73-85% continue to develop long-lasting side effects after therapy (Jensen et al., 2010). A major complication of radiation is salivary hypofunction, which is characterized by a partial or total loss of saliva accompanied with alterations in saliva composition. Consequently, the most common complaint in head and neck cancer patients who receive radiation therapy is xerostomia, a self-reported feeling of severe dry mouth (Dirix et al., 2006; Jensen et al., 2010).

Radiation-induced xerostomia and loss of saliva predispose individuals to develop dysphagia (difficulty to swallow), dysgeusia (loss of taste), periodontal and dental disease, chronic oral infections, and malnutrition. Not surprisingly the quality of life of these individuals is markedly reduced, and regrettably the available palliative options are short-lived and do not consistently improve quality of life (Dost and Farah, 2013;

Goldstein et al., 2014; Martinez Chibly, 2014; Nikles et al., 2013).

Newer radiotherapy modalities, such as Intensity modulated radiation therapy

(IMRT) and proton therapy (IMPT) allow a more precise delivery of radiation beams to the tumor while decreasing exposure of non-cancerous tissues. In animal studies as well as early clinical trials, IMRT improved quality of life by allowing partially sparing of the salivary glands from radiation (Kam et al., 2007; van Luijk et al., 2015; Nutting et al., 134

2011; Sio et al., 2016). Nevertheless, xerostomia and salivary hypofunction continue to be a persistent problem associated with radiation therapy, particularly in patients who are ineligible for sparing of the salivary glands due to the nature and anatomical localization of their tumor. Thus, it is of critical importance to develop regenerative therapies that restore saliva for patients receiving radiation therapy, as well as for those individuals currently living with loss of saliva and xerostomia. Regeneration of salivary glands following injury has been demonstrated by artificially ligating the main duct of the submandibular gland with a metal clip in rats (Carpenter et al., 2007; Osailan et al.,

2006). In these studies, ductal ligation led to progressive atrophy of acinar cells, but upon removal of the clip the remaining cells underwent a proliferative response that promoted glandular regeneration and recovery of saliva production. The molecular mechanisms that allow for glandular repair upon removal of the clip are still mostly unknown.

Adult tissues maintain their function in spite of being continuously exposed to damaging agents, due to the processes of homeostatic regeneration and wound healing.

These processes often require activation of a proliferation program that is initiated by signals received from the microenvironment, apoptotic, and inflammatory cells(Chen et al., 2003; Rakoff-Nahoum et al., 2004; Ryoo et al., 2004; Sato et al., 2011; Warner et al.,

2010). In some cases, the remaining differentiated cells proliferate to repair damage by undergoing dedifferentiation, self-duplication, or compensatory hypertrophy (Atanasoski et al., 2006; Aure et al., 2015; Nagy et al., 2001). But more frequently, it is the tissue- specific stem or progenitor cells that respond to these signals and undergo multiple rounds of cell division to replenish injured tissues(Bergmann and Steller, 2010; Blanpain 135 and Fuchs, 2014). Conventionally, adult stem and progenitor cells (SPCs) accomplish wound healing by dividing asymmetrically, making a copy of themselves while generating a differentiated cell, in order to maintain their numbers and support tissue function (Simons and Clevers, 2011). This choice of which cell fate decision to take – to expand the pool of stem cells or to differentiate into a more specialized cell – is regulated by complex interactions between adherens junctions, tight junctions, and apical-basal polarity complexes (Par-aPKC, Crumbs 3, and Scribble)(Lechler and Fuchs, 2005;

Martin-Belmonte and Perez-Moreno, 2011; Williams et al., 2011), as well as signals received from the stem cell niche (Hsu and Fuchs, 2012; Sato et al., 2011). Disruption of cell polarity, and specifically the Par3-aPKC complex, has been linked to ageing, tumorigenesis, and deficient wound healing (Mccaffrey et al., 2012; Niessen et al., 2013;

Vorhagen and Niessen, 2014; Xue et al., 2012). The Par3-aPKC complex is comprised of atypical protein kinase C (aPKC), partitioning defective 3 (PAR3) and 6 (PAR6), and cell division control 42 (CDC42), and is required for appropriate formation of the apical- lateral membrane borders (Goldstein and Macara, 2007; Henrique and Schweisguth,

2003). In addition, it has been studied extensively in the context of regulating cell division and cell fate in lower organisms and mammalian cells (Gómez et al., 2014;

Goulas et al., 2012; Niessen et al., 2013; Vorhagen and Niessen, 2014; Warner et al.,

2010).

Adult stem cells are attractive for designing new therapeutic approaches to start or potentiate wound healing in cases where it doesn’t normally occur, such as during cardiac disease, diabetes, neurodegeneration, and spinal cord injury (Kim et al., 2013; Mohsin et 136 al., 2011; Serakinci and Keith, 2006). In salivary glands, the use of adult stem cells has been proposed as an alternative treatment for xerostomia and salivary hypofunction (Patel and Hoffman, 2014; Pringle et al., 2013). Several research groups have identified multiple populations of SPCs in the developing salivary glands based on the expression of markers such as Keratin 5 (Ker5), Keratin 14 (Ker14), and c-kit, and studies have suggested their potential application to reestablish function in irradiated adult salivary glands by means of transplantation (Lombaert et al., 2008a; Ogawa et al., 2013). One question that remains unanswered is whether radiation has any direct effects on the stem cells that reside in the adult salivary glands, and whether these endogenous cells can be stimulated post-therapy to restore saliva. A drawback in answering these questions has been the lack of specific markers to delineate the diverse nature of salivary gland stem cells, particularly in the adult parotid gland, which is the most radiosensitive.

Our group previously identified a heterogeneous population of label-retaining cells (LRCs) in the salivary glands that comprises several stem and progenitor cell populations (Chibly et al., 2014). These cells survive radiation at least 30 days following treatment; however, function of the salivary glands is not restored. Understanding the mechanisms that prevent LRCs from repairing irradiated salivary glands will be instrumental in designing therapies to target specific populations of endogenous stem cells to promote wound healing without the need for transplantation. Here, we show that compensatory proliferation occurs in a fraction of LRCs in irradiated parotid salivary glands. Remarkably, this fraction is a unique population of LRCs that differs from

Keratin 5, Keratin 14, or c-Kit salivary gland progenitors, highlighting the importance of 137 further characterizing this population. Finally, we show that the increase in LRC proliferation is strongly associated with inactivation of aPKC and increased nuclear localization of the transcriptional coactivator YAP.

138

Methods

Mice and label-retaining assay

Experiments in this study were conducted in FVB, C57BL/6J, and Prkcz-/- mice, both male and female and repeated at least 2 times. Mice were maintained and treated in agreement with protocols approved by the University of Arizona Institutional Animal Care and Use Committee (IACUC). FVB, C57BL/6J, and Prkcz-/- mice were used in the present study and distributed to different experiments as indicated in the following sections.

Prkczfl/fl mice were generated in mixed background as described in Figure S3. The neomycin cassette was removed by crossing with mice carrying the FLP recombinase.

Prkczfl/fl mice were subsequently crossed with Sox2-Cre to generate Prkcz-/- mice. Sox2-Cre mice were obtained from Jackson Laboratory. At post-natal day 10, mice were given four intraperitoneal BrdU (5-Bromo-2’-deoxyuridine, Roche, Mannheim, Germany) injections at a dose of 3mg per 100g of body weight, or EdU (5-ethynyl-2’-deoxyuridine, Thermo

Fisher Scientific, Waltham, MA) injections at a dose of 10mg/100g of body weight 12 hours apart. At 10 weeks of age, mice were anesthetized via an intraperitoneal injection with Avertin (240mg/kg, Sigma, St Louis, MO) and euthanized by exsanguination for collection of the salivary glands.

Radiation treatment

Mice were anesthetized via intramuscular injection with ketamine/xylazine (50 mg/kg/10 mg/ml) prior to radiation treatment. A single dose of 5 Gy was targeted to the head and neck using a 60Cobalt Teletherapy instrument from Atomic Energy of Canada Ltd 139

Theratron-80, while the rest of the body was shielded from the radiation with >6 mm thick lead to avoid systemic effects. Radiation dosage calculations and maintenance of the cobalt source are conducted by the Experimental Radiation Shared Service of the Arizona Cancer

Center.

Immunohistochemistry and Immunofluorescence staining

Parotid, submandibular, and sublingual salivary glands were collected and immediately fixed in 10% neutral buffered formalin (Sigma) for 24 hours, then transferred to 70% ethanol and embedded in paraffin. Tissues were cut to 4μm thickness by the

Histology Service Laboratory in the Department of Cell Biology and Anatomy at the

University of Arizona and IDEXX Laboratories. Immediately before staining, tissue sections were baked at 37oC for 20 minutes to fully adhere tissues to glass slides. Tissues were then rehydrated in Histo-clear (National Diagnostics, Atlanta, GA), graded ethanol

(100%-50%) and distilled water, by performing 2 sequential washes in each solution for 5 minutes. Tissues were permeabilized in 0.2% TritonX (MP Biomedicals, Santa Ana, CA)

0.05% Tween20 (Thermo Fisher Scientific), in 1X PBS for 15 minutes prior to antigen retrieval, which was performed placing slides in 1mM citric acid buffer (pH 6.8) and boiling in microwave twice for 5 minutes. Slides were left in citric acid buffer for additional

20 minutes to cool down. From this point forward slides were washed three times for 5 minutes in fresh 1X PBS between steps. Non-specific binding sites were blocked with

300l of 0.5% NEN (Perkin Elmer, Waltham MA) followed by EdU staining as indicated by the manufacturer’s protocol (Click-iT® Plus EdU Alexa Fluor® 488 Imaging Kit, 140

Thermo Fisher Scientific). Tissues were incubated in primary antibody diluted in 1% BSA

(Sigma) overnight at 4oC. The following day, slides were incubated in secondary antibody diluted in 1% BSA for 1 hour at room temperature. Finally, tissues were counterstained with DAPI (1g/mL) and mounted with a solution of ProLong® Diamond Antifade

Mountant (Thermo Fisher Scientific). Slides were stored at 4 oC overnight before imaging with a Leica DM5500 microscope (Leica Microsystems, Wetzlar, Germany) and an

ORCA-Flash4.0 LT Digital CMOS camera (Hamamatsu Photonics K.K., Japan). We used primary antibodies anti-phospho-aPKCzT560 (1:250, abcam, Cambridge, UK. ab62372),

Par3 (1:50, Proteintech, Rosemont, IL. #11085-1-AP), Keratin 14 (1:400, Covance,

Princeton, NJ. PRB-155P), Keratin 5 (1:400, Covance, PRB-160P). Antibodies anti-Ki67

(1:200, #12202), c-kit (1:100, #3074), Cleaved caspase 3 (1:250, #9661), E-cadherin

(1:250, #3195) and Yap1 (1:250, #14074) were purchased from Cell Signaling (Danvers,

MA). Analysis for each marker was performed by manually counting positive cells from a minimum of 10 fields of view per slide using a 40X objective. A minimum of three slides

(three mice) per group were used for each marker. During analysis, we deemed the ductal compartment as everything that could be identifiable as a duct based on morphological features only, such as a rounded structure, the presence of a lumen, and tight cell-cell contacts. This compartment includes the excretory and striated ducts, as well as some of the intercalated ducts. Thus, the acinar compartment comprises all the remaining cell types in the salivary epithelium: acinar and myoepithelial cells, as well as intercalated ducts that based on morphology could not be identifiable as ducts. Acinar and ducal compartments 141 were analyzed individually and statistics were performed as described in the statistical analysis section.

Saliva collection

Stimulated saliva was collected from a minimum of 10 C57BL/6J and Prkcz-/- mice per group at 3 days following radiation treatment. Mice were given a 0.25 mg/kg dose of carbachol (Sigma-Aldrich) immediately before collection. During collection, mice were restrained with one hand and saliva was collected in pre-weighted tubes via vacuum aspiration. Saliva samples were immediately placed in dry ice and further stored at -80oC.

Saliva flow per minute was calculated for analysis and normalized to wild type unirradiated values.

Data analysis

Statistical analysis and graphing was performed using Graph-Pad software (version

7.01, La Jolla, CA). Cell counts from immunofluorescence stains were analyzed by a one- way analysis of variance (ANOVA), followed by Tukey’s multiple comparisons test. RT-

PCR data is normalized to GAPDH loading controls and expressed as fold-change vs wild type levels for each gene. Salivary flow rates were normalized to unirradiated wild type controls and analyzed by one-way ANOVA followed by Tukey’s multiple comparisons test.

142

Results

Radiation induces compensatory proliferation in a unique population of parotid acinar label-retaining cells

Using a BrdU or EdU label-retaining assay (Figure 3.1A), we previously demonstrated the presence of progenitor cells within a heterogeneous population of label- retaining cells in the murine salivary glands (Chibly et al., 2014). These cells are present

30 days following radiation treatment yet salivary function is not restored. We hypothesized that radiation impairs the ability of LRCs to initiate wound healing to replenish the injured salivary epithelium. It has been previously reported that in rat parotid and murine submandibular glands, radiation induces compensatory proliferation

6-9 days post-treatment (Bralic et al., 2005; Grundmann et al., 2010), presumably to attempt to restore function, but it is not clear whether SPCs participate in this response.

To determine whether LRCs contribute to the radiation-induced compensatory proliferation program, parotid tissue sections were immunostained for the proliferation marker Ki67 in combination with a marker for the LRCs (Figure 3.1B-C). Ki67-positive

LRCs were quantified within the acinar-enriched or ductal compartments at days 2 to 7 after radiation treatment (Figure 3.1A, D, Figure 3S.1A, See methods for description of each compartment).

Proliferation levels in unirradiated LRCs from the acinar-enriched compartment

(herein referred to as acinar LRCs) were 1.58±0.34%, and were significantly elevated at days 5 and 7 after radiation treatment (9.93±1.806% and 8.14±1.618%, respectively. 143

Figure 3.1D). Moreover, significantly higher levels of proliferation were also detected in non-LRCs in the acinar-enriched compartment at days 5 and 7 post-radiation

(9.873±1.837% and 8.555±1.435%, respectively) in comparison with untreated controls

(1.683±0.327%, Figure 3S.1B). Proliferation of acinar LRCs at day 4 post-radiation was not different than untreated acinar LRCs (data not shown), indicating that compensatory proliferation of acinar LRCs started at day 5. In contrast, ductal LRCs did not proliferate

(data not shown), and proliferation of non-LRCs in the ductal compartment was not different from untreated controls at the evaluated time points (Figure 3S.1C). Since initiation of compensatory proliferation did not involve activity of the ductal compartment, further analysis was focused exclusively on acinar LRCs.

Salivary gland LRCs comprise Ker5-positive (Ker5+), Ker14+, and c-kit+ progenitors, in addition to an unidentified population (Chibly et al., 2014). Because compensatory proliferation occurred only in a small fraction of the acinar LRCs, parotid glands of irradiated EdU-labeled mice were immunostained for Ki67 in combination with

Ker5, Ker14, or c-kit, to determine if these were the actively dividing sub-population of

LRCs following radiation (Figure 3.2). While 9.93% of acinar LRCs were dividing

(Ki67+) at day 5 after radiation treatment (Figure 3.1D), the fractions of proliferating c- kit+, Ker5+, and Ker14+ progenitors were only 0.97%, 1.12%, and 0.69%, respectively

(Figure 3.2D), indicating that the majority of the acinar LRCs that underwent compensatory proliferation are a unique population, which remains to be characterized. 144

Figure 3. 1: Acinar LRCs undergo compensatory proliferation 5-7 days post-radiation. A) Label-retaining assay: 10-day old FVB mice received four EdU (100mg/kg) or BrdU (30mg/kg) injections 12-hours apart. At 9 weeks of age, mice received a single 5Gy dose of radiation to the head and neck and parotid glands were collected at days 2-7 following treatment for analysis. Representative images of control and irradiated parotid glands immunostained for EdU (Green) and Ki67 (Magenta) are shown in panels B and C, respectively. Proliferative LRCs are shown with a white arrow. D) Percentage of Ki67- positive LRCs in acinar compartment. Statistical differences vs untreated group are represented by a star (p<0.05 by one-way ANOVA with follow up Tukey’s multiple comparisons test).

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Figure 3. 2: Cell populations undergoing compensatory proliferation in acinar compartment of the parotid gland. A-C) Representative images of irradiated FVB parotid glands stained for Ki67 (Green) and Keratin 5, Keratin 14, and c-kit (Magenta). White arrows point at dual positive (proliferating) cells. D) Percentage of proliferating cells (Ki67+) at day 5 post-radiation.

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T560 phosphorylation of aPKC is decreased in salivary gland LRCs following radiation

In optimal conditions, the Par-aPKC complex regulates proliferation of SPCs

(Goulas et al., 2012; Llado et al., 2015; Martin-Belmonte and Perez-Moreno, 2011;

Niessen et al., 2013; Vorhagen and Niessen, 2014). In order to understand why compensatory proliferation fails to promote regeneration in salivary glands, we aimed to determine how this process is regulated in acinar LRCs.

To first evaluate the positioning of the aPKC-Par complex within the LRCs, sections of parotid gland were immunostained for the polarity proteins Par3 and the active form of aPKC(referred to as aPKCT560), which contains a phosphorylated threonine residue at position 560 (Hirai, 2003) (Figure 3.3A-D). Par3 and aPKCT560 were found to be localized to the apical membrane of acinar LRCs at all the evaluated time points (Figure 3.3A-D). Interestingly, quantification of aPKCT560 within acinar

LRCs showed a decrease in the percentage of aPKCT560-positive acinar LRCs at days 5 and 7 following radiation, but not at day 4 (Figure 3.3E). The levels of aPKCT560 in ductal LRCs did not change across treatments (Figure 3.3F). As a reference, additional sections were stained for E-cadherin to visualize the lateral membranes. As expected, E- cadherin was found at the membrane of the LRCs and its localization did not change after radiation treatment (Figure 3S.2).

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Figure 3. 3: T560phosphorylation of aPKC in acinar LRCs decreases after radiation. A-B) Immunostaining of FVB mouse parotid cryosections for Par3 (Magenta) and EdU (Green). Areas delineated with a dashed line are shown in close-up images. C-D) Immunostaining of FVB mouse parotid FFPE glands for aPKCT560 (Magenta) and EdU (Green). Areas delineated with a dashed line are shown in close-up images, and white arrows point at dual positive cells. aPKCT560-negative LRCs are shown with a yellow star. E) Quantification of the percentage of aPKCT560-positive acinar LRCs. F) Quantification of the percentage of aPKCT560-positive ductal LRCs. Star represents statistical differences vs untreated group (p<0.05 by one-way ANOVA with follow up Tukey’s multiple comparisons test).

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Loss of aPKC induces a hyper-proliferative phenotype in acinar LRCs

Radiation-induced compensatory proliferation of acinar LRCs occurred at the same time as a decrease in aPKCT560 was observed, and in ductal LRCs, which did not proliferate, aPKCT560 was unchanged. Thus, we hypothesized that aPKC inhibits compensatory proliferation in acinar LRCs. In line with this hypothesis, proliferation of acinar LRCs was measured in a knockout mouse model for the Prkcz gene (Prkcz-/-,

Figure 3.4A-E, Figure 3S.3A, B) to test whether the loss of aPKC is sufficient to induce proliferation of acinar LRCs.

Analysis of unirradiated Prkcz-/- glands showed a statistically significant 6.56-fold increase in proliferation of the acinar LRCs when compared to C57BL/6J wild type controls (Figure 3.4E). To evaluate if loss of aPKC had an impact on the initiation of compensatory proliferation of acinar LRCs following radiation, the presence of Ki67 within acinar LRCs was quantified in wild type and Prkcz-/- mice 5 days post-radiation

(Figure 3.4C-E). Consistent with previous experiments, there was a 14.31-fold increase in proliferation of wild type acinar LRCs (WTLRCs) at day 5 post-treatment. Intriguingly, proliferation of irradiated Prkcz-/-- acinar LRCs (KOLRCs) at day 5 post-radiation was only 2.33-fold higher when compared to their respective unirradiated controls (Figure

3.4E). Taken together, these data suggest that aPKC inhibits proliferation of acinar

LRCs during glandular homeostasis and that radiation-induced compensatory proliferation is regulated by aPKC-dependent and aPKC-independent mechanisms. 149

Figure 3. 4: Initiation of radiation-induced compensatory proliferation in acinar LRCs is regulated by aPKC-dependent and aPKC-independent mechanisms. Immunostaining for Ki67 (magenta) and EdU (green) of parotid glands from wild type (C57BL/6J) and Prkcz-/- mice are shown in panels A-D. Areas delineated with a dotted line are shown in close-up panels (A’-D’). White arrows point at dual positive (proliferating) acinar LRCs. E) Quantification of the 150

percentage of Ki67-positive acinar LRCs. One-way ANOVA followed by Tukey’s multiple comparisons test was performed. Statistical differences between groups are shown with different letters (p<0.05). Same letters indicate non-statistically significant differences between groups. Acute radiation-induced loss of saliva occurs independently of aPKC

The higher baseline proliferation rates observed in untreated KOLRCs could potentially increase their sensitivity to radiation making them susceptible to cell death.

To test this, parotid glands from EdU-labeled mice were collected 24 hours following radiation treatment and stained for cleaved-Caspase 3 (Figure 3S.3C-F). This time point was chosen based on previous data from our lab showing higher levels of apoptosis in the acinar compartment, followed by acute loss of function after 3 days (Avila et al., 2009).

Consistent with previous research (Chibly et al., 2014), cleaved caspase 3 was not found in acinar WTLRCs (Figure 3S.3A, C). It was also undetectable in untreated KOLRCs

(not shown). In contrast, apoptosis was detected in a small subset of KOLRCs 24 hours post-radiation (Figure 3S.3B, D: White arrow).

Acinar cell death is thought to be responsible for acute loss of salivary function following radiation. To examine if the loss of aPKC was involved in acute loss of saliva post-radiation, saliva was collected from untreated and irradiated wild type (C57BL/6J) and Prkcz-/- mice, 3 days following radiation. In wild type animals, there was a reduction in saliva output in males and females by 22.51% and 22.71%, respectively. Similarly,

Prkcz-/- mice suffered a statistically significant decrease in saliva output 3 days following radiation by 27.56% and 30.22% for males and females, respectively (Figure 3.5).

However, there were no statistically significant differences between irradiated wild type and irradiated Prkcz-/- mice. 151

In combination, these data suggest that aPKC does not significantly contribute to the acute loss of saliva observed following radiation, which is consistent with our observation that levels of aPKCT560 within acinar LRCs were unchanged at day 4 following radiation. However, loss of aPKCresulted in elevated apoptosis following radiation, which suggests that aPKC may protect acinar LRCs from apoptosis upon radiation injury.

Radiation and loss of aPKCpromote nuclear localization of yap1 in acinar

LRCs

aPKC is well known for its role in maintaining cell polarity, and loss of polarity has been linked to increased proliferation (Llado et al., 2015; Mccaffrey et al., 2012;

Vorhagen and Niessen, 2014; Warner et al., 2010; Xue et al., 2012). Much less is known about how aPKC regulates the initiation of compensatory proliferation after injury, and what cellular pathways are regulated by or interact with aPKC to modulate proliferation.

The transcription coactivator Yap is an attractive target of study as a potential regulator of the initiation of compensatory proliferation in acinar LRCs, as it has been reported to have opposite roles than aPKC in regulating stem cell function (Llado et al., 2015).

Histological sections of parotid glands from irradiated and control EdU-labeled wild type mice were immunostained for Yap and EdU (Figure 3.6A, C), and the percentage of Yap- positive (Yap+) acinar LRCs were counted. In unirradiated controls 26.62% of acinar

LRCs were Yap+ (Figure 3.6E), and although a small increase was detected at day 5 following radiation to 37.94%, it was not statistically significant (p=0.22, Figure 3.6C, 152

E). Interestingly however, Yap was predominantly cytoplasmic in untreated acinar

WTLRCs (Figure 3.6A, A’, asterisk), with only 14.60% of Yap+ acinar LRCs having

Yap in the nucleus (Figure 3.6F). In contrast, the percentage of nuclear Yap+ acinar

WTLRCs was higher 5 days following radiation with 47.91% (p<0.001) of Yap+ acinar

WTLRCs having nuclear Yap (Figure 3.6C’: white arrow, Figure 3.6F), suggesting that radiation may promote transcriptional activity of Yap in acinar LRCs.

Next, Yap+ acinar KOLRCs were counted to determine if nuclear translocation of

Yap occurs downstream of aPKC (Figure 3.6B, D). The percentage of Yap+ acinar

KOLRCs in unirradiated mice was not statistically different from wild type controls

(23.39%, p=0.93, Figure 3.6 B, E), and it remained unchanged 5 days after radiation

(26.44%, p=0.99, Figure 3.6 D-E). However, nuclear Yap was highly elevated in unirradiated acinar KOLRCs to 49.26% (p<0.001, Figure 3.6B’: White arrows, Figure

3.6F), compared to WTLRCs, and persisted 5 days after radiation (50.10%, p<0.001,

Figure D’: white arrows, Figure 3.6F). Remarkably, the percentage of nuclear Yap in untreated acinar KOLRCs was similar to the percentage of nuclear Yap in irradiated

WTLRCs (p>0.90, Figure 3.6F), suggesting that aPKC acts upstream of Yap inhibiting

Yap’s ability to shuttle to the nucleus in acinar LRCs after radiation. Of note, there were a number of Ki67+Yap+ acinar LRCs (Figure 3S.4), which is suggestive of Yap playing a role in promoting proliferation of acinar LRCs.

153

Figure 3. 5: Acute loss of saliva following radiation is aPKC-independent. Saliva was collected from wild type (C57BL/6J) and Prkcz-/- mice as described in the methods section. Values for each mouse strain were normalized to their respective untreated control groups. Statistical differences between groups are represented by a star (p<0.05, One-way ANOVA followed by Tukey’s multiple comparisons test).

154

Figure 3. 6: Radiation induces nuclear translocation of Yap in acinar LRCs. Immunostaining for Yap (magenta) and EdU (green) of parotid glands from wild type (C57BL/6J) and Prkcz-/- mice are shown in panels A-D. Areas delineated with a dotted line are shown in close-up panels (A’-D’). White arrows point at Yap-positive LRCs with nuclear Yap. E) Quantification of the percentage of Yap-positive acinar LRCs. F) Quantification of the percentage of Nuclear-Yap+ LRCs in acinar compartment (white arrows). One-way ANOVA followed by Tukey’s multiple comparisons test was performed. Starts represent statistical differences vs wild type untreated group (p<0.05).

155

Discussion

It has been a matter of debate whether salivary gland SPCs contribute to maintain homeostasis in the adult glands, and whether they promote repair following damage.

Here, we report that a unique population of label-retaining cells (LRCs) in the acinar compartment of the parotid gland initiate compensatory proliferation in response to radiation-induced injury. LRCs are a heterogeneous population of salivary gland progenitor cells (Chibly et al., 2014); thus, our results support the idea that salivary SPCs participate in the response to radiation injury.

In regeneration models of salivary glands using the ductal ligation technique, most acinar cells undergo apoptosis, yet the gland is able to regenerate from the surviving cells (Carpenter et al., 2007; Osailan et al., 2006). The extensive acinar cell death observed in these studies suggested that regeneration likely occurred from expansion and differentiation of a different cell type, presumably residing in the ductal system.

Similarly, pioneering studies using 3H-Thymidine labeling assays concluded that the intercalated ducts replenished the acinar cells by a differentiation process (Denny et al.,

1993; Man et al., 2001). It was speculated that salivary gland stem cells reside in the ducts, and that these cells are responsible for maintaining homeostasis in the adult salivary glands (See (Pringle et al., 2013) for review). A more recent study challenged this point of view as it demonstrated that Mist1-positive acinar cells, which are thought to be fully differentiated, contribute to glandular growth and homeostasis through a process of self-duplication. The authors propose that such process may be the primary mechanism 156 for maintenance of homeostasis in the adult salivary glands and may also contribute to regeneration of the gland in ductal ligation experiments (Aure et al., 2015). In the aforementioned studies it is evident that proliferation is a key process during salivary gland regeneration, and the authors point to the possibility that multiple cell types participate in this proliferative process to orchestrate tissue repair. Our data supports a model in which salivary gland progenitors participate in the proliferative response that results from radiation; however, it also suggests that known ductal progenitors do not contribute to replenish the acinar compartment during the initial response to radiation injury. Additionally, our study shows that LRCs in the acinar compartment – which includes acini and intercalated ducts – have proliferative potential, and may contribute to glandular homeostasis and repair (Aure et al., 2015). It is well known, however, that radiation-induced loss of salivary function does not improve overtime (Jensen et al.,

2010). This result underscores the importance of elucidating which specific cell populations do and do not participate in the radiation-induced compensatory proliferation program and understanding how this process is regulated.

We found that Ker5, Ker14, and c-kit progenitors do not significantly participate in the initiation of radiation-induced compensatory proliferation in salivary glands.

Instead, a unique population of acinar LRCs responds more readily at this stage.

Although it is unknown whether acinar LRCs express Mist1, we previously reported that salivary acinar LRCs are not fully differentiated as they did not stain positive for - amylase in vivo (Chibly et al., 2014). Thus, we speculate that the acinar cell populations responsible for repair after radiation injury (possibly acinar LRCs) differ from those 157 undergoing self-duplication during homeostasis, and we propose the existence of a unique population of acinar progenitor cells with the ability to proliferate in response to radiation. Further studies will be necessary to fully characterize this population.

Radiation-induced compensatory proliferation has been described in other systems and appears to have regulatory mechanisms that are conserved across species

(Llado et al., 2015; Warner and Longmore, 2010; Zimmerman et al., 2013). For instance, in the Drosophila larval wing imaginal discs, radiation induces apoptosis and disrupts epithelial cell polarity by targeting the Par-aPKC polarity complex. In turn, disruption of cell polarity promotes proliferation of the surviving cells (Warner et al., 2010). Similarly, high levels of proliferation are observed in the stem cell compartment of the mouse intestine in response to radiation, which also allows for intestinal regeneration (Hua et al.,

2012; Llado et al., 2015; Metcalfe et al., 2014). Moreover, the Par-aPKC complex regulates cell division of stem cells in the Drosophila midgut (Goulas et al., 2012), and deletion of aPKC in Lgr5+ stem cells in the mouse intestine potentiates proliferation of this population and accelerates regeneration through increased Yap and -Catenin activity (Llado et al., 2015). In the present study, radiation induced nuclear translocation of Yap in acinar LRCs and decreased the percentage of acinar LRCs with aPKCT560

(Figure 3E), which we interpreted as inactivation of the kinase. Both events were first detected at day 5 following radiation when compensatory proliferation of acinar LRCs initiated, and after acute loss of function was observed. In addition, genetic ablation of aPKC resulted in increased proliferation and nuclear localization of Yap in acinar LRCs, suggesting that Yap acts downstream of aPKC. We propose that, similar to its function 158 in mouse intestine, aPKC inhibits proliferation of acinar LRCs in salivary glands through regulation of Yap’s localization. Interestingly, the combination of radiation and loss of aPKC resulted in greater levels of proliferation than with loss of aPKCalone, which could indicate that proliferation of acinar LRCs is only partially inhibited by aPKC and that additional mechanisms exist to regulate radiation-induced compensatory proliferation in this population. Additionally, the deletion of aPKC did not potentiate radiation-induced compensatory proliferation of acinar LRCs (Figure 4E: comparison between irradiated wild type vs irradiated Prkcz-/-), as opposed to how Lgr5+ stem cells in the intestine respond to radiation (Metcalfe et al., 2014). One possibility is that acinar

LRCs reach their maximum proliferative potential after radiation injury through a combination of inactivation of aPKC and additional unidentified mechanisms, in which case the loss of aPKC would only account for the fraction of elevated proliferation caused by inactivation of aPKC in wild type animals, and not for the unknown mechanisms.

Based on our findings, we propose a model in which radiation inactivates aPKC by reducing its phosphorylation in a subpopulation of salivary progenitors, promoting nuclear translocation of Yap and activating a proliferation program. Future studies are necessary to determine how compensatory proliferation in salivary progenitors can be modulated to avoid or reverse chronic loss of salivary function following radiation.

159

Supplementary material

Figure 3S. 1 Acinar and ductal compartments of the gland were analyzed individually as described in the methods section. Ductal compartment is shown in the delineated areas with a dotted line in panel A). All remaining cells were considered part of the acinar compartment. B) Quantification of the percentage of Ki67+ (proliferative) cells in the acinar compartment. C) Quantification of the percentage of Ki67+ cells in the ductal compartment. Star represents statistical differences vs the Untreated group (p<0.05) by one-way ANOVA followed by Tukey's multiple comparisons test.

160

Figure 3S. 2 Immunostaining of FVB parotid glands for E-cadherin (magenta) and EdU (green). Delineated areas areas are shown in close-up panels. E-cadherin is localized at the lateral membranes in untreated (A) and Irradiated (B) samples

161

Figure 3S. 3 A) Cloning strategy for the generation of Prkcz-floxed mice. (B) Identification of the floxed allele by southern blotting. Prkczfl/fl mice were subsequently crossed with Sox2-Cre to generate Prkcz-/- mice. Immunostaining of parotid glands from wild type (C57BL/6J) and Prkcz-/- mice for EdU and cleaved-Caspase 3 is shown in panels C-D. Close-up images of delineated areas are shown in E-F. White arrow points at an apoptotic LRC.

162

Figure 3S. 4 Immunostaining of irradiated FVB mouse parotid glands for EdU (green), Ki67 (magenta) and Yap (Gray). White arrows point at Ki67+Yap+ LRCs.

163

APENDIX III. APKC-DEPENDENT INHIBITION OF NUCLEAR YAP MEDIATES RESTORATION OF SALIVARY FUNCTION WITH IGF-1

Funding support: NIH R01DE023534 to KHL. AMC was supported by CONACyT, Mexico.

Key words: Salivary glands, label retaining cells, IGF-1, radiation, progenitors, head and neck cancer, aPKC, Yap1.

Conflict of interest: the authors declare none

164

Introduction

Loss of saliva and xerostomia are common side effects in individuals with head and neck cancer treated with radiotherapy. Although there is no cure available for these conditions, salivary function has been partially restored in animal studies by means of transplantation (Lombaert et al., 2008a; Ogawa et al., 2013), gene therapy (Delporte et al., 1997; Shan et al., 2005), and pharmacological intervention (Grundmann et al., 2010;

Hill et al., 2014; Xiao et al., 2014). In humans, a phase I clinical trial using adenoviral- based delivery of aquaporin 1 gene (AdhAQP1) showed improvements in saliva production in five out of eleven patients (Baum et al., 2012). In addition, regeneration of salivary glands using stem cell-based approaches has been suggested as a potential therapy for radiation-induced loss of saliva and xerostomia (Pringle et al., 2013).

Pharmacological stimulation of endogenous salivary stem and progenitor cells to promote regeneration has been previously reported. In mice, administration of Glial cell line–derived Neurotrophic Factor (GDNF) 1-day post-radiation rescued the submandibular gland from radiation-induced damage. The authors of this study concluded that recovery is likely to occur through the expansion of the Lin-CD24+c-

Kit+Sca1+ subpopulation of progenitor cells in the submandibular gland, in which the receptor for GDNF is enriched (Xiao et al., 2014). Neurturin (NRTN), another member of the GDNF-family of neurotrophic factors, was also shown to promote epithelial regeneration in irradiated submandibular explants from mouse embryos (Knox et al.,

2013). Regeneration was likely due to the protective effect of neurturin on the 165 parasympathetic nerves, which in turn promotes survival of Keratin-5 progenitors (Knox et al., 2010). It is important to note that both GDNF and NRTN were administered during early acute stages of the glandular response to radiation injury. In the case of GDNF, which was administered in vivo, this corresponds to a stage in which chronic loss of function has not yet developed. This is problematic when targeting specific cell populations, such as Lin-CD24+c-Kit+Sca1+, because the chronic effect of radiation upon these populations is still unknown, and whether they can be stimulated at chronic stages remains to be elucidated. Finally, previous work from our lab showed that systemic administration of Insulin-like growth factor 1 (IGF-1) at 4 to 7 days post-radiation effectively restores saliva production in mice (Grundmann et al., 2010); however, whether IGF-1-induced recovery of function involves stimulation of endogenous salivary stem and progenitor cells has not been studied.

Acinar label-retaining cells (LRCs), which are comprised of multiple types of salivary progenitors, are present long-term after radiation and undergo compensatory proliferation in response to radiation injury (Chibly et al., 2014, Chapter III), suggesting that there is potential to target this population to promote regeneration. Stem and progenitor cells normally undergo compensatory proliferation to promote tissue regeneration following injury (Bergmann and Steller, 2010; Blanpain and Fuchs, 2014); however, the pathways involved in entry and exit of this proliferative program, and how the process is regulated in salivary glands, are not completely understood. In acinar LRCs of the parotid salivary glands, initiation of radiation-induced compensatory proliferation involves a decrease in phosphorylation of atypical protein kinase C zeta (aPKC) and 166 elevation of nuclear Yes-associated protein (Yap) levels (Chapter III). Interestingly, IGF-

1 has been shown to induce activation of aPKC (Escribano, 2003; Liu et al., 1998), and aPKC is a known regulator of Yap during tissue regeneration (Llado et al., 2015); thus, determining whether these pathways are regulated by IGF-1 during regeneration of salivary glands may be instrumental for the development of new therapies for salivary gland dysfunction and xerostomia.

Not much is known about the functions of Yap in the salivary glands. In general,

Yap and Taz (transcriptional co-activator with a PDZ-binding domain, also known as

WW domain containing transcription regulator 1, or WWTR1) are the main effectors downstream of the Hippo pathway, which is associated with orchestrating the balance between apoptosis, proliferation, and differentiation, to ensure proper organ formation without enabling overgrowth (Varelas et al., 2014; Yu and Guan, 2013). Yap and Taz do not have DNA-binding domains but elicit transcription activation via cognate transcription factors, such as SMADs, p63, RUNX, and PAX, which greatly diversifies the transcriptional output of the pathway (Varelas et al., 2014). Additionally, it is known to participate in tissue regeneration following injury, and defects in this pathway have been associated with tumorigenesis (Varelas et al., 2014). In the embryonic submandibular gland, increased levels of Yap and its paralog Taz, were associated with assembly of tight junctions during the process of branching, which suggested a role of

Hippo pathway in establishing cell polarity during development (Enger et al., 2013). 167

Here, we show that IGF-1 promotes T560-phosphorylation of aPKC in acinar

LRCs of irradiated salivary glands, and that aPKC is an upstream regulator of Yap in acinar LRCs. Finally, we propose that radiation leads to hyperactivation of Yap in acinar

LRCs in an aPKC-dependent manner, and recovery of salivary function with IGF-1 requires a proper balance in levels of aPKCT560 and nuclear Yap.

168

Methods

Mice and label-retaining assay

Prkczfl/fl mice were generated in mixed background as described in Figure S3. The neomycin cassette was removed by crossing with mice carrying the FLP recombinase.

Prkczfl/fl mice were subsequently crossed with Sox2-Cre to generate Prkcz-/- mice. Sox2-Cre mice were obtained from Jackson Laboratory. At post-natal day 10, mice were given four intraperitoneal EdU (5-ethynyl-2’-deoxyuridine, Thermo Fisher Scientific, Waltham, MA) injections at a dose of 10mg/100g of body weight 12 hours apart. At 10 weeks of age, mice were anesthetized via an intraperitoneal injection with Avertin (240mg/kg, Sigma, St

Louis, MO) and euthanized by exsanguination for collection of the salivary glands.

Radiation treatment

Theratron-80, while the rest of the body was shielded from the radiation with >6 mm thick lead to avoid systemic effects. Radiation dosage calculations and maintenance of the cobalt 169 source are conducted by the Experimental Radiation Shared Service of the Arizona Cancer

Center. For “Day 30” treatments, mice were irradiated at 6 weeks of age. All other mice that received radiation were treated 5 or 8 days prior to collection of the glands.

IGF-1 Injections

Mice were given a maximum of four IGF-1 doses (5g/mouse) via tail-vein injections 24-hours apart at days 4 to 7 after radiation treatment. For “Day 5” treatments, mice received only one injection at day 4 post-radiation and salivary glands were harvested

24 hours later. For All other time points mice were given a total of 4 injections, and tissues were collected at the time points indicated in each experiment.

Immunohistochemistry and Immunofluorescence staining

Histology Service Laboratory in the Department of Cell Biology and Anatomy at the

(100%-50%) and distilled water, by performing 2 sequential washes in each solution for 5 minutes. Tissues were permeabilized in 0.2% TritonX (MP Biomedicals, Santa Ana, CA)

0.05% Tween20 (Thermo Fisher Scientific), in 1X PBS for 15 minutes prior to antigen retrieval, which was performed placing slides in 1mM citric acid buffer (pH 6.8) and 170 boiling in microwave twice for 5 minutes. Slides were left in citric acid buffer for additional

20 minutes to cool down. From this point forward slides were washed three times for 5 minutes in fresh 1X PBS between steps. Non-specific binding sites were blocked with

300l of 0.5% NEN (Perkin Elmer, Waltham MA) followed by EdU staining as indicated by the manufacturer’s protocol (Click-iT® Plus EdU Alexa Fluor® 488 Imaging Kit,

Thermo Fisher Scientific). Tissues were incubated in primary antibody diluted in 1% BSA

Mountant (Thermo Fisher Scientific). Slides were stored at 4 oC overnight before imaging with a Leica DM5500 microscope (Leica Microsystems, Wetzlar, Germany) and an

ORCA-Flash4.0 LT Digital CMOS camera (Hamamatsu Photonics K.K., Japan). We used primary antibodies anti-phospho-aPKCzT560 (1:250, abcam, Cambridge, UK. ab62372), anti Yap (1:250, Cell Signaling, Danvers, MA. #14074), and phospho-AKTSer473 (Cell signaling, #4060). Analysis for each marker was performed by manually counting positive cells from a minimum of 10 fields of view per slide using a 40X objective. A minimum of three slides (three mice) per group were used for each marker. During analysis, we deemed the ductal compartment as everything that could be identifiable as a duct based on morphological features only, such as a rounded structure, the presence of a lumen, and tight cell-cell contacts. This compartment includes the excretory and striated ducts, as well as some of the intercalated ducts. Thus, the acinar compartment comprises all the remaining cell types in the salivary epithelium: acinar and myoepithelial cells, as well as intercalated 171 ducts that based on morphology could not be identifiable as ducts. Acinar and ducal compartments were analyzed individually.

Saliva collection

Stimulated saliva was collected from C57BL/6J and Prkcz-/- mice at 30 days after radiation treatment. Mice were given a 0.25 mg/kg dose of carbachol (Sigma-Aldrich) immediately before collection. During collection, mice were restrained with one hand and saliva was collected in pre-weighted tubes via vacuum aspiration. Saliva samples were immediately placed in dry ice and further stored at -80oC. Saliva flow per minute was calculated for analysis and normalized to wild type unirradiated values.

Data analysis

Statistical analysis and graphing was performed using Graph-Pad software (version

7.01, La Jolla, CA). Cell counts from immunofluorescence stains were analyzed by a one- way analysis of variance (ANOVA), followed by Tukey’s multiple comparisons test.

Salivary flow rates were normalized to unirradiated wild type controls and analyzed by one-way ANOVA followed by Tukey’s multiple comparisons test.

172

Results

Decreased T560phosphorylation of aPKC in acinar LRCs is associated with chronic loss of function.

The unrepaired acute radiation damage to the salivary epithelium is thought to predispose the glands to chronic loss of function. Previous data from our lab showed that radiation caused a decrease in the active phosphorylated form of aPKC (aPKCT560) in acinar LRCs 5-7 days after treatment. In turn, genetic ablation of Prkcz suggested that aPKC negatively regulates the initiation of compensatory proliferation induced by radiation in acinar LRCs. Thus, we aimed to determine if aPKC in acinar LRCs contributes to chronic loss of salivary function following radiation.

To this end, parotid glands were harvested 30 days after radiation treatment

(Figure 4.1 A), and were processed for analysis by immunofluorescence staining of aPKCT560 and EdU as a marker of the LRCs (Figure 4.1B-E). Analysis showed a statistically significant decrease in the percentage of aPKCT560-positive acinar LRCs 30 days following radiation (Figure 4.1F). Consistent with a previous report, aPKCT560 in ductal LRCs did not change after radiation (data not shown). Next, to measure the direct impact of Prkcz loss in salivary function, stimulated saliva production was measured in irradiated Prkcz-/- and wild type (C57BL/6J) mice. Saliva output of irradiated wild type mice was reduced to 76% of untreated values, while in Prkcz-/- mice radiation caused a greater decrease in saliva production to 65% of control values (Figure 4.1G). Notably, 173 there was no change in salivary output of untreated Prkcz-/- mice compared to wild type controls.

Systemic post-therapeutic administration of IGF-1 partially restores salivary function in mice 30 days post-therapy, and full recovery of function is observed at 60 and

90 days after treatment (Grundmann et al., 2010); however, the mechanisms that govern salivary gland regeneration upon administration of IGF-1 are unknown. Therefore,

T560phosphorylation of aPKC in acinar LRCs was evaluated in irradiated mice treated with IGF-1 to establish whether aPKC in salivary progenitors is associated with recovery of function (Figure 4.2A-C). The percentage of aPKCT560-positive acinar LRCs was higher 30 days post-radiation in mice that received the IGF-1 injections versus the ones that didn’t (Figure 4.2D). Moreover, saliva output in IGF-1 treated Prkcz-/- mice was significantly reduced compared to controls 30 days after radiation (Figure 4.2E), indicating that IGF-1 fails to restore salivary function in the absence of aPKC.

In combination, these data show a strong negative association between

T560phosphorylation of aPKC in acinar LRCs and chronic loss of saliva, and indicates that aPKC is required for restoration of salivary function with IGF-1 in mice.

174

Figure 4. 1: Decrease in aPKCT560 in acinar LRCs is associated with chronic loss of function. A) 10-day old C57BL/6J mice were labeled with EdU as described in the methods section. At 6 weeks of age, mice were irradiated with a single 5 Gy dose of radiation to the head and neck and parotid glands were harvested for analysis 30 days later (at 10 weeks of age). B-E) Parotid glands were immunostained for EdU and phosphorylated aPKCT560. Bottom right corner of C and E are zoomed in images of the areas delineated with a dotted line. White arrows point at acinar LRCs with positive staining for aPKCT560. F) Quantification of the percentage of aPKCT560-positive acinar LRCs in untreated and irradiated animals. Star represents statistically significant differences (p<0.05 unpaired 2-sided t-test) G) Carbachol-stimulated saliva was collected from untreated and irradiated C57BL/6J and Prkcz-/- mice. Statistical differences are represented with a star (One-way ANOVA followed by Tukeys multiple comparisons test). 175

Figure 4. 2: Recovery of salivary function with IGF-1 requires aPKC. A) 10-day old C57BL/6J mice were labeled with EdU as described in the methods section. At 6 weeks of age, mice were irradiated with a single 5 Gy dose of radiation to the head and neck. At 4-7 days post- radiation, mice received tail vein injections with IGF-1 (5g/mouse) 24-hours apart. Parotid glands were harvested for analysis 30 days later (at 10 weeks of age). B-C) Parotid glands from IGF-treated mice were immunostained for EdU and phosphorylated aPKCT560. Bottom right corner of C is a zoomed-in image of the area delineated with a dotted line. D) Quantification of the percentage of aPKCT560-positive acinar LRCs in IGF-treated mice. Values of untreated and irradiated animals are shown with a black and blue dotted lines, respectively. E) Carbachol-stimulated saliva was collected from wild type (C57BL/6J) untreated and IGF-treated Prkcz-/- mice. Statistical differences are represented with a star (p<0.05 unpaired 2-sided t-test).

176

IGF-1 regulates nuclear levels of Yap in acinar LRCs in an aPKC- dependent fashion

Generally, IGF-1 signals preferentially via the AKT pathway to promote survival (Siddle,

2011). In addition, aPKC has been linked with both IGF-1 and PI3K-AKT pathways

(Chou et al., 1998). Thus, to evaluate the potential contribution of AKT signaling in acinar LRCs to recovery of salivary function with IGF-1 administration, parotid glands from irradiated and control mice were stained for the active phosphorylated form of AKT

(AKTSer473). IGF-1 was administered via tail vein injections at days 4-7 post-radiation.

Activation of AKT was evaluated 24 hours following the first and last IGF-1 doses (days

5 and 8 post-radiation) (Figure 4.3A). There were no changes in the percentage of

AKTSer473-positive acinar LRCs in irradiated animals when compared to untreated controls (Figure 4S.1A-B, D). Notably, there was a transient decrease in the percentage of AKTSer473-positive acinar LRCs at day 5 post-radiation upon administration of the first dose of IGF-1, but it was not sustained by day 8 post-treatment (Figure 4S.1C-D).

We previously showed that nuclear translocation of Yap is tightly associated with the initiation of compensatory proliferation in acinar LRCs following radiation treatment

(Chapter III). In addition, it has been reported that nuclear Yap is necessary for the regenerative process elicited by the Lgr5+ stem cells in the intestine (Cai et al., 2010).

Therefore, modulation of the hippo pathway may be important during salivary gland regeneration induced by IGF-1. To address this question, wild type mice (C57BL/6J) received IGF-1 at days 4-7 post-radiation and parotid glands were harvested 24 hours 177 after the first and last IGF-1 injections (Figure 4.3A). Consistent with previous research

(Chaper III), quantification of the percentage of Yap-positive acinar LRCs (Yap+LRCs) was not different across treatments (data not shown); however, the percentage of

Yap+LRCs with nuclear Yap ([nuclear]Yap+LRCs) was elevated in irradiated samples

(Figure 4B,D; white arrows). IGF-1 administration induced a decrease in the percentage of Yap4.+LRCs with nuclear Yap when compared to irradiated mice that did not receive

IGF-1 (Figure 4.3C-D). Notably, levels of Yap+LRCs with nuclear Yap following systemic administration of IGF-1 were still elevated in comparison with unirradiated controls (Figure 4.3D, dotted red line). Further, to evaluate the significance of nuclear

Yap at later stages of regeneration, parotid glands from irradiated and IGF-1-treated mice were analyzed 30 days after treatment with radiation (Figure 4.2A). In irradiated animals, levels of nuclear Yap in acinar LRCs remained elevated, and administration of IGF-1 reduced the percentage of Yap+LRCs with nuclear Yap compared to irradiated animals

(Figure 4.4A-C).

aPKC has been reported to inhibit Yap during intestinal regeneration via direct phosphorylation of S109 and T110. Thus, we hypothesized that IGF-1 regulates nuclear translocation of Yap in acinar LRCs in an aPKC-dependent manner. In line with this hypothesis, parotid glands from IGF-treated and irradiated Prkcz-/- mice were immunostained for Yap and EdU. Whereas the percentage of Yap+LRCs with nuclear

Yap was significantly lower in IGF-treated wild-type mice compared to irradiated animals (Figure 4.4D, F: compare with dotted red line), IGF-1 failed to replicate this response in Prkcz-/- mice (Figure 4.4F). 178

Figure 4. 3: IGF-1 reduces levels of nuclear Yap in irradiated acinar LRCs. A) 10-day old C57BL/6J mice were labeled with EdU as described in the methods section. At 9 weeks of age, mice were irradiated with a single 5 Gy dose of radiation to the head and neck. At 4-7 days post- radiation, mice received tail vein injections with IGF-1 (5g/mouse) 24-hours apart. Parotid glands were harvested for analysis at 5 and 8 days post-radiation. B-C) Parotid glands from irradiated and IGF-treated mice were immunostained for EdU and Yap. Bottom panels in B and C are zoomed-in images of the areas delineated with a dotted line. D) Quantification of the percentage of [nuclear]Yap-positive acinar LRCs in irradiated and IGF-treated mice. Percentage of [nuclear]Yap-positive acinar LRCs in untreated animals is shown with a red dotted line for reference.

179

Figure 4. 4: IGF-1 inhibition of nuclear Yap is mediated by aPKC. A-B) Parotid glands from irradiated and IGF-treated mice, collected at 30 days post-radiation, were immunostained for EdU and Yap. Bottom panels are zoomed-in images of the areas delineated with a dotted line. C) Quantification of the percentage of [nuclear]Yap-positive acinar LRCs in irradiated and IGF-treated mice. Percentage of [nuclear]Yap-positive acinar LRCs in untreated animals is shown with a red dotted line for reference. D-E) Parotid glands from IGF-treated wild type and Prkcz-/- mice, collected at 5 days post-radiation, were immunostained for EdU and Yap. Bottom panels are zoomed-in images of the areas delineated with a dotted line. F) Quantification of the percentage of [nuclear]Yap-positive acinar LRCs in wild type and Prkcz-/- IGF-treated mice. Percentage of [nuclear]Yap-positive acinar LRCs in irradiated animals that did not receive IGF-1 is shown with a red dotted line for reference.

180

Discussion

The use of stem cells has been widely proposed as a potential treatment for radiation-induced salivary hypofunction and xerostomia primarily by means of transplantation (Lombaert et al., 2008a, 2011; Ogawa et al., 2013; Patel and Hoffman,

2014; Pringle et al., 2013; Yoo et al., 2014). Alternatively, our study suggests that pharmacological stimulation of the endogenous pool of salivary LRCs with IGF-1 may promote regeneration without the need for transplantation. However, systemic administration of growth factors, such as IGF-1, prompts concern of developing de novo malignancies (Chae et al., 2015; Ogilvy-Stuart and Gleeson, 2004). For this reason, their utilization as treatment for radiation-induced salivary hypofunction may be contingent upon the development of a delivery mechanism that allows targeted administration to the salivary glands (i.e. gene therapy or nanoparticles). Alternatively, elucidating the mechanisms that drive salivary regeneration upon administration of IGF-1 may be used for the development of safer and more efficient therapies. Here we show that recovery of salivary function with IGF-1 is mediated by modulating the levels of nuclear Yes- associated protein (Yap) in acinar LRCs in an aPKC-dependent manner, and independently of AKT signaling.

We previously showed that acinar LRCs, which comprise multiple salivary gland progenitors, are present 30 days post-radiation, when chronic loss of salivary function is evident (Chibly et al., 2014; Grundmann et al., 2010). Here, IGF-1 was administered after acute loss of function developed, and during a stage in which acinar LRCs initiate a 181 compensatory proliferation program (Chapter III). This might suggest the existence of a therapeutic window in which salivary progenitors can be effectively targeted to rescue the glands from radiation-induced damage. Numerous pathways are likely involved in promoting salivary regeneration with IGF-1; however, the aPKC and Hippo pathways are of particular interest because of recent observations demonstrating that radiation inhibits aPKC and promotes nuclear translocation of Yap in acinar LRCs during the initiation of compensatory proliferation (Chapter III). Presumably, these mechanisms converge with secretory pathways, as administration of IGF induced the generation of amylase-secreting acinar cells in irradiated glands (Grundmann et al., 2010).

A functional role of aPKC during epithelial regeneration has been demonstrated in the intestine, where genetic ablation of aPKC improved intestinal regeneration after radiation, by increasing the proliferative activity of Lgr5+ stem cells through enhanced- catenin and Yap signaling (Llado et al., 2015). In contrast with observations in intestinal stem cells, genetic deletion of Prkcz exacerbated radiation-induced loss of saliva, and recovery of salivary function with IGF-1 involved an increase in aPKC phosphorylation in acinar LRCs. These differences might point at tissue-specific functions of aPKC during epithelial regeneration. Nonetheless, our findings suggest that nuclear translocation of Yap is a common downstream effector of aPKC signaling in stem and progenitor cells during radiation-induced injury and subsequent regeneration.

Multiple miRNAs, adaptor proteins, and G-coupled receptors, have emerged as potential regulators of Yap function (Mo et al., 2014; Varelas et al., 2014). Other studies 182 have also suggested that activation of Yap is downstream of cell polarity regulators, including the aPKC-Par complex (Grzeschik et al., 2010; Llado et al., 2015; Sun and

Irvine, 2011; Wang et al., 2016, Chapter III). In support of our observations, genetic deletion of aPKC in Lgr5+ stem cells promotes activation of Yap function during intestinal regeneration following radiation damage (Llado et al., 2015). Contrasting studies in Drosophila eye and wing epithelium showed that expression of a membrane- bound form of aPKC (aPKCCAAX) induced activation of Yorkie (Drosophila orthologue of

Yap) (Grzeschik et al., 2010; Sun and Irvine, 2011). We previously reported that aPKC inhibits nuclear translocation of Yap in acinar LRCs of the salivary glands, as nuclear

Yap was elevated in acinar LRCs of Prkcz-/- mice. Here, we show that radiation injury to the salivary epithelium promotes an increase in nuclear translocation of Yap in acinar

LRCs, which persists up to 30 days following injury. Post-therapeutic administration of

IGF-1 reduces the levels of nuclear Yap in acinar LRCs in comparison with irradiated animals that did not receive IGF-1. Moreover, IGF-1 failed to decrease levels of nuclear

Yap in acinar LRCs of irradiated Prkcz-/- mice, which indicates that translocation of Yap to the nucleus in acinar LRCs during salivary gland regeneration is also dependent on aPKC

It is important to note that levels of nuclear Yap did not return to control levels 30 days after treatment, when administration of IGF-1 has been shown to restore salivary output (Grundmann et al., 2010). Some possible explanations are that nuclear translocation of Yap is more effectively repressed before injury, or that Yap is somewhat dispensable for salivary function during homeostasis. Consistent with the latter, studies in 183 the intestine have shown that conditional knockout of Yap using a Villin-Cre system did not have any obvious defects in differentiation, proliferation, cell death, or migration along the crypt-villus axis during development (Barry et al., 2013; Cai et al., 2010).

However, intestinal regeneration was severely impaired in mice with Yap-deficient intestines following injury with dextran sodium sulfate (DSS) or radiation (Barry et al.,

2013; Cai et al., 2010). Very interestingly, loss of Yap promoted a decrease in crypt proliferation upon DSS-induced injury (Cai et al., 2010), but induced crypt hyperplasia in the irradiated intestines (Barry et al., 2013). Although these remarks point to seemingly opposing roles of Yap in regulating stem cell function, another study demonstrated that genetic ablation of Yap specifically in the Lgr5+ stem cells led to deficient crypt regeneration following radiation (Gregorieff et al., 2015), further supporting that Yap is required for proper stem-cell driven regeneration. On the other hand, hyperactivation of

Yap has also been associated with tumorigenesis and improper regeneration (Cai et al.,

2010; Camargo et al., 2007; Zhou et al., 2011). In combination, these observations might explain our results in the salivary glands if we consider that Yap is likely to exert different functions depending on its cellular localization, as well as the influence from upstream regulators such as aPKC. Indeed, our data suggests that a delicate balance in levels of aPKC and nuclear Yap is necessary to achieve regeneration of the salivary glands following radiation injury.

In addition to regulating proliferation of intestinal stem cells, Yap’s association with specific transcription factors (i.e. Klf4) induces differentiation into secretory goblet cells (Imajo et al., 2015). In parallel, IGF administration reduced compensatory 184 proliferation and promoted differentiation into amylase-secreting acinar cells in irradiated glands (Grundmann et al., 2010); thus, is it feasible that regulation of nuclear Yap by

IGF1 is associated with differentiation of the LRCs into functional acinar cells, contributing to the recovery of saliva secretions.

We propose that during salivary gland homeostasis, nuclear Yap in acinar LRCs is more effectively repressed by aPKC and other upstream regulators. After radiation injury, lower levels of aPKCT560 result in hyperactivation of Yap and overall glandular dysfunction. Finally, IGF-1 normalizes the levels of aPKCT560 while maintaining sufficient levels of nuclear Yap to promote regeneration. Future studies will be necessary to determine whether aPKC and Yap, or their downstream effectors, can be targeted directly in salivary progenitors to promote salivary gland regeneration.

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Supplementary Material

Figure 4S. 1: IGF-1 does not activate AKT in irradiated acinar LRCs. A-C) Parotid glands from untreated, irradiated, and IGF-treated C57BL/6J mice were immunostained for EdU and phosphorylated AKTSer473. D) Quantification of the percentage of AKTSer473-positive acinar LRCs in untreated, irradiated, and IGF-treated mice.