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Student Research Submissions

Spring 5-7-2021

Proton Pump Inhibitors Promote Apoptosis in Jurkat T Proton Pump Inhibitors Promote Apoptosis in Jurkat T

Lymphocytes Lymphocytes

Ashley Utz

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Recommended Citation Recommended Citation Utz, Ashley, "Proton Pump Inhibitors Promote Apoptosis in Jurkat T Lymphocytes" (2021). Student Research Submissions. 386. https://scholar.umw.edu/student_research/386

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ABSTRACT

Cancer cells are known to rely on the anaerobic energy pathway of glycolysis even under normoxic conditions, resulting in measurable intracellular acidification that may trigger cell death by apoptosis. In normal cells, the pH is restored by activation of voltage-gated proton pumps, preventing apoptosis. Proton pump inhibitors (PPIs), such as omeprazole, inhibit the action of these voltage-gated proton pumps. Research has shown that omeprazole is also capable of inducing caspase-dependent apoptosis in Jurkat T-lymphocytes, but this area of study remains largely unexplored. The goal of this research was to determine the temporal dynamics of caspase activity in Jurkat cells treated with omeprazole, dexlansoprazole, or esomeprazole. Caspase activation was observed with fluorescence microscopy and the fluorogenic caspase probe, L-bisaspartic acid rhodamine 110 (D2R). All three PPIs were shown to induce significant caspase activity when incubated in the presence of the drug over a 30-hour period, with dexlansoprazole showing comparative activity to the positive control, doxorubicin. Microfluidic devices were utilized to capture cells for real-time analysis, but only the doxorubicin showed caspase activity during these trials, with an onset time of 3.66 ± 1.67 hours. The microfluidic device experiments showed a decrease in cell-to-device binding affinity following the incubation period, indicating that PPI exposure alters the cell surface receptors in some capacity. Future studies will focus on addressing these challenges to allow for the elucidation of the intensity and timing of caspase activation, which will be beneficial for evaluating PPIs as potential cancer therapeutics.

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ACKNOWLEDGMENTS

I would first like to express my sincerest gratitude to Mrs. Irene Piscopo Rodgers for her generous funding through the Irene Piscopo Rodgers ’59 and James D. Rodgers Student Research Fellowship I. The funding from this fellowship made this project possible, particularly amidst the difficulties brought about by the pandemic. I would also like to give my deepest thanks to my research advisor, Dr. Randall Reif, for his unwavering support of my academic endeavors. During our first ever lab session for introductory chemistry, I stayed behind to ask Dr. Reif about the possibility of pursuing research with his undergraduate team. That turned out to be one of the best decisions I have ever made, alongside my decision to attend UMW, and I will forever be grateful for the chance to do research on this team and to learn from Dr. Reif.

I would also like to personally thank Dr. Kelli Slunt, Dr. Leanna Giancarlo, and Dr. Davis Oldham for their willingness to serve on my committee. During my time at UMW, I was fortunate to have the chance to take classes with these amazing professors, and their support and guidance has been invaluable. I extend my sincere thanks to the Department of Chemistry and Physics and the Department of Biology; these departments, which have both become like a home to me in my time here, have graciously provided the resources, space, and time necessary for this project to be completed.

I am indebted to the other members of the Reif URES team, who have all contributed to

my research endeavors over the past three years. I want to especially thank Shreya Murali for her assistance in data collection and processing, as well as Ramsey Cotton and Sofia Bondarenko for their help in fabricating the microfluidic devices.

To my family, I want to express my most sincere appreciation for their steadfast support

of my college pursuits. I want to thank my parents, Kenneth Utz and Pamela Utz, for always encouraging me to do my best, and for reminding me to take some time for myself when I have needed it. I thank my grandfather, Gibson Harrell, who guided me in becoming the person I am today and helped make it possible for me to attend college. On the most disheartening of days, I know he is always still cheering me on and inspiring me to not give up.

I want to thank Kylie Sunamoto, my roommate and a dear friend, for her companionship

and her jokes as I sat on the floor in the early hours of the morning, typing away at this thesis. Last but certainly not least, I want to express my heartfelt gratitude and appreciation for my partner, Bình Dương, for her patience and support as I navigated this research. Her willingness to talk through the toughest of questions with me until I came up with a plan of action, her continuous encouragement in the form of routine pep talks, and her infallible ability to make me laugh have all been indispensable to both my health and the accomplishment of my goals. I am most fortunate to be able to dedicate to and share this research with those I love the most.

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LIST OF ABBREVIATIONS

BSA- bovine serum albumin

BSL- biosafety level

CAD- caspase-activated DNase

CIPN- chemotherapy-induced peripheral neuropathy

D2R- fluorogenic caspase probe (aspartic acid)

DMSO- dimethyl sulfoxide

DNA- deoxyribonucleic acid

FBS- fetal bovine serum

ICAD- inhibitor of caspase-activated DNase

MPT- mitochondrial permeability transition

PARP- poly-ADP-ribose polymerase

PBS- phosphate buffered saline

PDMS- polydimethylsiloxane

PEG- polyethylene glycol

PPI- proton pump inhibitor

R110- rhodamine 110

RPMI- Roswell Park Memorial Institute

TCR- T-cell receptor

TfR- transferrin receptor

TABLE OF CONTENTS

ABSTRACT……………………………………………………………………...……………..ii

ACKNOWLEDGEMENTS……………………………………………...….………………iii

LIST OF ABBREVIATIONS…………………………………………...…………..………iv

LIST OF FIGURES……………………………………………………….…………..……….2

1. INTRODUCTION……………………………………………...………………………….3 1.1 Cancer and its Survival Mechanisms………………………………………….………….3

1.2 Apoptosis and the Caspases………………………………………………………………5

1.3 Proton Pump Inhibitors………………………………………………...…………………9

1.4 Temporal Dynamics of Caspase Activation…………………………………………….15

2. MATERIALS AND METHODS…………………………….…………….…………..20 2.1 Microfluidic Device Fabrication…………………………………………………………20

2.2 Coating of Microfluidic Devices with Antibody Ladder……………………...…………20

2.3 Preparation of PPIs, Doxorubicin, and Fluorogenic Probe…………………...………….21

2.4 Cell Culture………………………………………………………………………..……..22

2.5 Baseline Caspase Activation Study……………………………………………….……..22

2.6 Temporal Detection of Caspase Activity………………………………..……………….23

2.7 Binding Affinity Study……………………………………………………….………….25

2.8 Analysis of Images……………………………………………………………....……….26

2.9 Safety Considerations…………………………………………..………………………..27

3. RESULTS………………………………………………………………….………………..28 3.1 Baseline Caspase Study………………………………………………………………….28

3.2 Microfluidic Device Temporal Trials…………………………………...……………….30

3.3 Binding Affinity Study…………………………………………………………………..32

4. DISCUSSION………………………………………………………………….…………..35

5. CONCLUSION………………………………………………………………………….....50

REFERENCES……………………………………………………………………………..…53

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LIST OF FIGURES

FIGURE 1: Extrusion of Protons by a Typical Cancer Cell

FIGURE 2: Caspase Cascade of Mitochondria-Dependent Apoptosis

FIGURE 3: Morphological Features of Apoptosis

FIGURE 4: Structures of Proton Pump Inhibitors

FIGURE 5: Mechanism of Activation of Proton Pump Inhibitors

FIGURE 6: Microscope Setup and Anatomy of Microfluidic Devices

FIGURE 7: Microfluidic Device Antibody Ladder

FIGURE 8: Caspase Cleaving Fluorescent Probe

FIGURE 9: Percentage of Apoptotic Cells by Hour for Each Drug

FIGURE 10: Fluorescence Images from the Baseline Activation Study

FIGURE 11: Fluorescence Images from the Microfluidic Device Temporal Trials

FIGURE 12: Percentage of Bound Cells by Hour for Each Drug

FIGURE 13: White-Light Pre-wash and Post-Wash Images for Each Drug

FIGURE 14: Fluorescence Pre-Wash and Post-Wash Images for Each Drug

FIGURE 15: Optimal Cell Color and Confluency

FIGURE 16: Residual PDMS Polymer Stuck in the Microfluidic Device Capillary

FIGURE 17: Influx of Bubbles to the Microfluidic Device Observation Area

FIGURE 18: Bubble in Field of Vision of a Fluorescence Image

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1. INTRODUCTION

1.1 Cancer and its Survival Mechanisms

In 2021, 608,570 cancer deaths are predicted to occur in the United States (Siegel, 2021).

Although treatment options for cancer have been widely expanded over the last century, many

human solid tumors remain troublesome to treat. Major problem areas of existing therapies

include general systemic toxicity associated with drugs, acquired resistance to chemotherapeutic

drugs (multi-drug resistance), and disruption of drug efficacy in acidic environments (Fais,

2010). Cancer cells favor increased glucose uptake and fermentation of glucose to lactate, even

when in the presence of ample oxygen (Liberti, 2016). This phenomenon, known as the Warburg

Effect, works in conjunction with regional hypoxia and nutrient deficiency to result in a

measurable acidification of the interior of the cancer cells (Damaghi, 2013). Cancer cells

proliferate optimally when they exhibit a slightly alkaline pH, around 7.0-7.6, and will begin to

undergo metabolic stress if the intracellular pH falls below 6.7 (Persi et al., 2018). Thus, if left

unregulated, this intracellular acidification caused by the Warburg Effect results in the induction

of apoptosis (Gottlieb et al., 1996; Sharma et al., 2015; and Lagadic-Gossmann et al., 2004).

To avoid this fate, cancer cells maintain survivable pH levels via the overexpression and

upregulated activation of plasma membrane proteins, including pumps and transporters, to

facilitate the release of H+ ions to raise the alkalinity of the cell’s interior (Huber et al., 2010).

These pathways create a reverse pH gradient compared to normal cells, in which the cancer cell’s

interior is more alkaline than its environment (Persi et al., 2018). The cells release lactate and H+

via monocarboxylate transporters (Gillies, 2002) and Na+-driven proton extrusion (Gallagher et

al., 2008), turning their extracellular environment acidic (Figure 1). When these pathways

malfunction or are subjected to inhibition via drugs, the efflux of H+ can no longer take place,

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resulting in the acidification of the cell’s intracellular environment, and eventually leading to cell

death. Further methods of pH regulation vary by cell type but may include carbonic anhydrases

(Barathova et al., 2008) or V-ATPase, a vacuolar-type H+/K+-ATPase (Hinton et al., 2009).

Figure 1: The extrusion of protons (shown in blue) performed by a typical cancer cell. The cell employs several methods of maintaining intracellular alkalinity, including sodium-driven proton extrusion, V-ATPase activity, and monocarboxylate transporters (MCT). The overexpression of these mechanisms leads to a measurable acidification of the extra-cellular environment (Huber et

al., 2010). Image created in BioRender.com.

V-ATPase has been shown to be particularly important for cancer cell survival, acting as

a promoter of proliferation and metastasis by facilitating the acidification of the extracellular

environment as it maintains the pH within the cell (von Schwarzenberg, 2013 and Wiedmann,

2012). In cancer, V-ATPase expression is upregulated to maintain a survivable pH within the cell

(Nelson and Harvey, 1999). Elevated expression of V-ATPase mRNA has been associated with

poor survival in ovarian cancer patients (Lee et al., 2015), suggesting that regulation of V-

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ATPase expression is essential for increasing the chances of patient survival. The inhibition of

V-ATPase has been shown to limit cancer proliferation and induce apoptosis in a variety of cell

lines (Whitton, 2018), as it leads to an increased expression of the pro-apoptotic tumor-

suppressor protein p53 (Williams et al., 1999 and Long et al., 1998). Since intracellular

acidification has been shown to precede caspase activation in cells undergoing mitochondria-

dependent apoptosis (Matsuyama, 2000), drugs which lead to intracellular acidification may be

viable for cancer therapies. These findings suggest that inhibition of V-ATPase is a valuable

target for anti-cancer research, particularly where the induction of apoptosis in tumor cells is

desired.

1.2 Apoptosis and the Caspases

Apoptosis, or programmed cell death, is characterized by distinct biochemical

mechanisms (Elmore, 2007). It is essential for normal cell turnover, maintenance of tissue

homeostasis, immune system regulation, and many other critical biological processes. Improper

regulation of apoptosis can lead to neurodegenerative diseases, ischemic damage, and

autoimmune disorders (Elmore, 2007), and is widely recognized as the most common cause of

cancer development (Finkel, 2001). Apoptosis proceeds via the extrinsic pathway, where the cell

responds to external signals, or the intrinsic pathway, which results from internal stimuli. The

role of PPIs in cancer cell death is currently attributed to the function of the intrinsic pathway.

The loss of apoptotic suppression seen in this pathway may be due to the absence of growth

factors, hormones, and cytokines that would typically inhibit the death of the cell (Elmore,

2007). Pro-intrinsic-apoptosis signals can also include radiation, toxins, hypoxia, hyperthermia,

infection by bacteria or viruses, and free radical oxidative stress (Susin et al., 2000).

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Pro-apoptotic signals result in changes to the inner mitochondrial membrane that cause

the cascade of apoptosis (Figure 2) to begin. These signals trigger the mitochondrial permeability

transition (MPT) pore to make a conformational change to the open position (Elmore, 2007).

This change results in the loss of the mitochondrial transmembrane potential and the release of

granzyme b into the inner membrane space, facilitating the movement of pro-apoptotic proteins

cytochrome c, Smac/DIABLO, and HtrA2/Omi into the cytosol, where they activate the caspase-

dependent mitochondrial pathway of apoptosis (Saelens et al., 2004; Cai et al., 1998; Du et al.,

200; Loo et al., 2002; Garrido et al., 2005).

Figure 2: The cascade of apoptosis, beginning with the example stimulus of severe DNA

damage. Pro-apoptotic signals, like the Bax protein, facilitate the release of cytochrome c from the mitochondria. Cytochrome c binds to APAF-1, forming the apoptosome complex, where the

procaspase-9 is cleaved to the active caspase-9 form. Caspase-9 activates the executioner caspase-3 which cleaves the death substrates and goes on to cleave other caspases to facilitate the amplification of the apoptotic signal (Saelens et al., 2004; Cai et al., 1998; Du et al., 2000; Loo

et al., 2002; Garrido et al., 2005). Image created in BioRender.com.

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Caspases, a family of cysteine proteases, cleave their substrates on the C-terminus of

aspartic acid residues to drive the signaling cascade of caspase-dependent apoptosis (Julien and

Wells, 2017). Once the pro-apoptotic proteins move to the cytosol, cytochrome c binds to the

protease adaptor APAF-1, and the complex binds to procaspase-9, forming the apoptosome and

leading to the activation of caspase-9, the initiator caspase for the intrinsic pathway (Chinnaiyan,

1999). Once caspase-9 is active, the full cascade effect of apoptosis begins, as active caspases

auto-catalytically cleave additional caspases, resulting in significant and exponential

amplification of the initial signal (Poreba, 2015). As such, apoptosis is an “all or nothing”

process; once the caspase cascade begins, it continues autonomously. Smac/DIABLO and

HtrA2/Omi inhibit the activity of proteins that normally suppress apoptosis, aiding the activation

of the apoptotic cascade (Loo et al., 2002).

In the execution phase of apoptosis, the effector caspases are activated (Elmore, 2007).

These “executioner” caspases activate cytoplasmic endonuclease and other proteases, leading to

the degradation of the nuclear material and nuclear and cytoskeletal proteins. Executioner

caspases resulting in the blebbing and budding of the plasma membrane and the orderly death of

the cell (Slee et al., 2001). Caspase-3 is recognized as the most important effector caspase, as it

cleaves the endonuclease inhibitor of caspase-activated DNase (ICAD) to release CAD, which

then degrades chromosomal DNA and leads to chromatin condensation (Sakahira et al., 1998).

Caspase-3 also triggers the formation of the apoptotic bodies and is responsible for the cleavage

of gelsolin, which results in disruption of the cytoskeleton, intracellular transport, cell division,

and signal transduction (Kothakota et al., 1997). The function of caspase-3, as well as the other

effector caspases and the initiator caspases that activate them, is essential for the occurrence of

apoptosis and the regulation of cell turnover in a manner that is not disruptive or damaging to the

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surrounding tissues.

A hallmark of apoptosis is the absence of an inflammatory reaction to cell death

(Kurosaka et al., 2003). As the apoptotic cell undergoes controlled death, the cellular membrane

experiences blebbing which is then followed by karyorrhexis and budding (Figure 3), where the

cell fragments separate into compact apoptotic bodies (Elmore, 2007). These apoptotic bodies

are quickly phagocytosed by macrophages, preventing secondary necrosis and eliminating the

production of anti-inflammatory cytokines (Savill and Fadok, 2000; Kurosaka et al., 2003). The

lack of inflammatory response and avoidance of necrosis makes apoptosis the most orderly and

least disruptive form of cell death. Unfortunately, cancer cells employ several regulatory

pathways to inhibit apoptosis (Shen, 2010), and upregulation of V-ATPase to maintain

intracellular alkalinity is just one method. Furthermore, many cancer cell lines show resistance to

the caspase-9 pathway of apoptosis (Reif, 2010). Drugs that restore normal levels of caspase-

dependent apoptosis represent promising treatment strategies (Wong, 2011). Drugs that return

expression of the V-ATPase to normal levels, or compensate for its overexpression, remain

viable targets for anti-cancer therapies.

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Figure 3: The cycle of physical symptoms of apoptosis exhibited by dying cells. The cellular

membrane begins to undergo blebbing, which is joined by karyorrhexis. Blebbing and fragmentation of the nuclear material continue to occur until the cell is broken up into small apoptotic fragments which are then engulfed by surrounding macrophages (Elmore, 2007).

Image created in BioRender.com.

1.3 Proton Pump Inhibitors

Proton pump inhibitors are a class of drugs with the specific purpose of inhibiting gastric

acid secretion by inhibiting gastric V-ATPase on the luminal surface of the parietal cell

membrane (Shin, 2010). Gastric V-ATPase is the enzyme responsible for secreting gastric acid

via an electroneutral, ATP-dependent hydrogen-potassium exchange mechanism, pumping K+

into the cell and H+ out of the cell (Sachs, 1976). Gastric V-ATPase has an identical function to

that observed in cancer cells exhibiting the Warburg Effect. This makes the PPIs, which inhibit

gastric V-ATPase, potential targets for inhibiting cancer V-ATPase activity.

PPIs are commonly utilized as a secondary or supportive treatment to cancer therapies, as

the acidity of cancer cells presents problems for targeted anti-tumor treatments, which are

commonly weak bases (Fais, 2010). These weak bases encounter the extracellular environment

of the tumor cell, which is acidic due to the cell’s pH-regulating process, and are protonated, at

which point they are no longer membrane-permeable (Fais, 2010). PPIs have been shown to

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inhibit V-ATPase pathways, resulting in intracellular acidification that makes the tumor cells

more susceptible to anticancer agents (Ikemura, 2017). The drugs are most often taken orally for

their traditional anti-acid applications, but may also be administered intravenously or via direct

injection, making them more bioavailable for proton pump inhibition at a faster rate (Wu et al.,

2019).

The pharmacophore of all PPI drugs is 2-pyridylmethylsulfinylbenzimidazole, and the

constituent R groups vary from drug-to-drug (Figure 4) (Roche, 2006). The chemical reactivity

and inhibitory potency of the drugs is heavily influenced by the electron-donating or electron-

withdrawing nature of the substituents (Roche, 2006). Upon initial application to the body, the

PPIs exist in a pro-drug or inactive form. To convert the inactive PPIs to their active, inhibitor

forms, both the pyridine and benzimidazole rings must be protonated according to their pKa

values (Shin and Kim, 2013 and Roche, 2006). The pyridine nitrogen is protonated first at a pKa

ranging from 3.83 to 4.53 (Figure 5) (Roche, 2006). In their traditional applications, this first

protonation occurs once the drug is already uptaken by the gastric parietal cells, and will render

the drug membrane-impermeable, so it cannot leave the cell. The second protonation of the

benzimidazole nitrogen follows immediately at a pKa ranging from 0.11 to 0.79 (Roche, 2006).

This protonation generates the monocation of the charged benzimidazole ring and the unionized

pyridine ring, which can conduct an intramolecular nucleophilic attack to produce sulfenamide

and sulfenic acid. These products form covalent disulfide bonds with the sulfhydryl groups on

the CYS813 or CYS822 residues in the V-ATPase enzyme, holding it in the inactive

conformation. This inhibition of V-ATPase is thus irreversible, and the acid-secretion ability of

V-ATPase is terminated (Roche, 2006). The body must synthesize new V-ATPase enzymes for

acid secretion to resume.

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The parietal cells the PPIs normally target have a low intracellular pH around 0.8,

meaning their intracellular environment is where protonation and the action of the PPIs takes

place (Roche, 2006). However, this is not the case for cancer cells, which exhibit an intracellular

pH that is alkaline. Instead, the protonation and activation of the PPIs is believed to occur in the

cancer cell’s acidic microenvironment (Lu et al., 2017). It remains unknown whether the cancer

cells can up-take the PPIs, or whether the drug’s inhibitory effects occur solely in the cells

immediate external environment. It is hypothesized that the PPIs are activated to form their

reactive sulfenamide and sulfenic acid compounds and still form disulfide bonds with the V-

ATPase’s cysteine residues on the extracellular portion of the transmembrane enzyme, but the

specific residues remain largely unexamined (Lu, et al., 2017).

Figure 4: The structures of the PPIs utilized in this study. The basic structure of the

benzimidazole and pyridine rings are consistent for each drug, but various constituent groups differ. The electron-donating or electron-withdrawing nature of these constituents influences

inhibitor potency. (Adapted from Morschel et al, 2018).

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Figure 5: The overview of the mechanism of activation of the PPIs. The first step is protonation of the pyridine nitrogen, followed by the protonation of the benzimidazole nitrogen. Two monocations of the protonated benzimidazole structure exist; one with an ionized pyridine

nitrogen and one with an unionized pyridine nitrogen. It is the monocation with the unionized monocation that can undergo an intramolecular nucleophilic attack to generate the active sulfenic

acid and sulfenamide products. These products readily form disulfide bonds with sulfhydryl groups in the cysteine residues of V-ATPase, and thus inhibit the enzyme’s acid-secretion

function. (Image taken from Shin and Kim, 2013).

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Despite the mechanism remaining largely unknown, the inhibiting effects of the PPIs

have shown increased sensitization of cancer cells to chemotherapeutics, translating to a

reduction in drug resistance (De Milito and Fais, 2005). Omeprazole and esomeprazole were

shown to enhance the effects of anticancer agents through inhibition of V-ATPase in human

melanoma, adenocarcinoma, and lymphoma (Luciani et al., 2004). Furthermore, omeprazole was

shown to induce apoptosis in Jurkat cells via cleavage of procaspase-3 and PARP (Scaringi,

2004). This is promising, since caspase-3 is the most important effector caspase, and it partakes

in many of the cleavage steps that directly result in membrane blebbing and chromatin

condensation. Although studies have shown that PPIs can induce cell death via a caspase-

independent pathway involving reactive oxygen species (Canitano et al., 2016 and De Milito et

al., 2007), current methods of treatment view PPIs as primarily supportive care. PPIs have

additional benefits as therapeutics, as they only become active in acidic environments, meaning

they can selectively target tumor cells, and they generally carry only mild side effects (Walsh,

2015). Existing chemotherapeutic agents are commonly cell-cycle-specific, and can only

eliminate cancer cells when they are actively dividing. This presents another problem with drug

resistance, as human tumors often contain cells that proliferate very slowly (Dey-Guha, 2011).

This leaves behind residual cancer cells that eventually divide and reform the tumor.

Chemotherapeutics which are non-cell-cycle-specific, such as doxorubicin, are generally

toxic to the system. These drugs often confer serious side-effects and are non-specific to

targeting cancer cells. Instead, the agent destroys all cells it encounters, leading to the damaging

of the patient’s tissues that can be permanent (Johnson-Arbor and Dubey, 2021). Among the

most common chemotherapy side effects are nausea and vomiting (Rapoport, 2017).

Manifestations of nausea and vomiting come in the acute form, which is reasonably treatable,

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and the delayed form, which is poorly manageable (Rapoport, 2017). Additional gastrointestinal

side-effects are commonly seen in chemotherapy, including oral and gastrointestinal mucositis

(Cinausero et al., 2017). Uncontrolled chemotherapy-induced diarrhea is also common, with

potentially fatal consequences of dehydration and electrolyte imbalance, and may require

reductions or cessations of treatment (Nurgali, 2018). Cognitive impairments may manifest or

persist long after the end of treatment (Castel et al., 2017). Chemotherapy-induced peripheral

neuropathy (CIPN) is also very common, and can cause a host of different symptoms (Nurgali,

2018). Long-term CIPN is associated with depression, ataxia, insomnia, and gastrointestinal

dysfunction (Kerckhove et al., 2017). While some advancements have been made in reducing the

severity or duration of these side-effects, they persist frequently in patients undergoing

chemotherapy. Doxorubicin, the positive control in this study, is capable of inducing

cardiomyopathy at any point following treatment. Doxorubicin-induced cardiomyopathy has no

cure and will persist throughout the duration of the patient’s life, eventually contributing to an

earlier mortality (Johnson-Arbor and Dubey, 2021).

Proton pump inhibitors are well-tolerated in clinical administrations, with serious adverse

events being extremely rare (Thomson, 2010). The most common concern reported with long-

term or acute PPI usage is rebound acid reflux upon the cessation of treatment. Long-term use of

PPIs has not been shown to cause carcinoid tumors or increase the risk of gastric cancers,

although some links are currently being investigated (Thomson, 2010). PPIs have been proven to

be safe and effective in managing gastrointestinal reflux disease, healing peptic ulcers, and

reducing gastritis. They have become one of the most-prescribed medications employed by

healthcare providers in modern medicine (Ambizas, 2017). These drugs are associated with an

increased risk of developing community-acquired pneumonia, which could be risky for

15

immunocompromised patients, such as those undergoing chemotherapy (Johnson, 2013), but are

generally well-tolerated by the body. The lack of reported adverse effects and the generally low

toxicity of PPI drugs make them not only safe for use as supportive therapy, but also affirm their

potential as standalone, low-toxicity therapeutic agents.

Since PPI drugs are non-cell-cycle-specific but are generally nontoxic to the normal cells

and tissues, and trigger both caspase-dependent and caspase-independent apoptosis, they may be

more effective in drug-resistant or apoptosis-resistant forms of cancer. Inducing the same levels

of death in cancerous cells while maintaining the viability of the normal tissues could bring forth

a breakthrough in lowering the toxicity of cancer treatments. However, little research has been

conducted to uncover the temporal dynamics of caspases and biochemical mechanisms within

apoptotic PPI pathways. Furthermore, the apoptotic capabilities of other commercially-available

PPI drugs, including new-generation PPIs like dexlansoprazole, have not yet been documented.

Thus, this study seeks to elucidate the apoptotic effects of select PPI drugs on Jurkat cells via

real-time monitoring of caspase activity.

1.4 Temporal Dynamics of Caspase Activation

Caspase activation and inhibition present major fields of study in cancer research.

Unfortunately, cancer cells are notorious for being resistant to apoptosis, and multidrug

resistance is rampant across cell lines (Fais, 2010). This presents problems for traditional

caspase-targeted therapies, and often renders them ineffective. In cases where the inhibition of

caspases is selected as a course of treatment, the cell will often die via caspase-independent

pathways, including the lysis of cells resulting from granule exocytosis (Kolenko, 2000).

Caspase-independent pathways of apoptosis have recently showed promise in emerging studies,

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yet present their own unique problems. Caspase-independent pathways are often pro-

inflammatory, resulting in the recruitment of inflammatory cytokines (Giampazolias, 2018).

While the triggering of the immune inflammatory response can be harmful in some cases,

(Ammirante et al., 2010), the host immune response can be utilized to aid the body’s response to

chemotherapy (Zitvogel et al., 2008).

Prior research has been conducted to examine the general timing of caspase activity in

living cells undergoing apoptosis as a natural event, but the temporal dynamics of caspases

remains largely unknown. Caspase-3 activation has been shown to occur seven hours post-

induction (Sundquist et al., 2006). Further data exists on caspases involved in the extrinsic

pathway, but very little is known about the timeline for caspase activation in the intrinsic

pathway. Furthermore, individual cellular assays could provide a more robust outline of caspase

activation than multi-cell studies. Microfluidic devices designed by Reif et al. in 2009 (Figure 6)

provide an affinity surface for cell capture, while maintaining gas and waste exchange and

allowing for a continuous medium supply. Cells may be bound to the affinity surface via

antibodies, which have been utilized extensively in cell affinity chromatography (Pappas, 2007).

The Reif et al. device utilizes an affinity ladder of biotinylated bovine serum albumin (BSA),

NeutrAvidin, and anti-CD71 to hold cells stationary for observation. When the cells pass through

the microfluidic channel, those expressing the cell surface receptors corresponding to anti-CD71

are retained in the affinity column. The antibody ladder (Figure 7), which is applied according to

a procedure by Kuida et al., is carefully selected to contain an antibody that does not induce

apoptosis (Reif, 2009). Once cells are bound to the affinity channel, they remain viable for up to

ninety-six hours (Liu, 2008), allowing for long periods of observation.

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Figure 6: The microscope setup with a zoom-in of the Reif et al. microfluidic device. The device

features a PDMS polymer housing with a molded microfluidic channel for cell capture. For experimentation, the device is connected to a Y-junction of Teflon tubing, which is then attached

to syringes of PBS and FBS-PBS on syringe pumps. Image created in BioRender.com.

Figure 7: The antibody ladder utilized for cell capture. Biotinylated BSA is applied to the microscope slide. The NeutrAvidin is then applied, which binds to the BSA. The detection

antibody, anti-CD71 is then applied to the NeutrAvidin layer. Image created in BioRender.com.

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Apoptosis can be observed in captured cells utilizing fluorescence microscopy and

fluorogenic probes. Rhodamine 110 (R110) is a naturally fluorescent, membrane-impermeable

molecule known as a fluorophore (Reif et al., 2010). Conjugation of specific tetrapeptide

sequences to the R110 molecule renders it non-fluorescent and membrane-permeable, allowing it

to be utilized as a fluorogenic caspase probe. Cells uptake the conjugated molecule, (Asp)2-

rhodamine 110 (D2R), where its covalently-bound aspartic acid residues, are susceptible to

cleavage by active caspases. Active caspases recognize and cleave the aspartic acid tetrapeptide

substituents on the non-fluorescent R110 molecule, freeing R110 and converting it to its mono-

substituted form, which fluoresces when excited with 488 nm LED light (Martinez, 2010). The

fluorescent molecule is released upon cleavage and remains within the cell, allowing for the

detection of caspase activity (Figure 8). Once the aspartic acid residues are cleaved, the free

R110 molecule is again membrane-permeable, but will leave the cell at a very slow rate. Thus,

when real-time observation is conducted, the amount of fluorescence detected in a D2R-treated

cell correlates directly to the amount of caspase activation occurring in the cell (Martinez, 2010).

These methods, when combined with the use of affinity surfaces for observation of the cells,

allow for the measurement of caspase activity in response to drug treatment over a period of

several hours and on an individual-cell basis.

19

Figure 8: The process of a caspase cleaving the aspartic acid residues from the D2R molecule.

Once cleaved, the R110 fluorophore returns to its native fluorescent form. Since D2R is absorbed into the cytosol of the cell, the cleaved R110 fluorophore is retained for a period, and the cancer

cell will fluoresce upon excitation with 488 nm light. Image created in BioRender.com.

In this work, various PPI drugs were evaluated for their ability to induce caspase-

dependent apoptosis in Jurkat T lymphocytes. Induction of mitochondrial apoptosis was verified

utilizing cell affinity capture and fluorescence microscopy. Once drugs were shown to induce

apoptosis, temporal dynamics assays were performed to determine the timeframe for general

caspase activity. Preliminary microscopy studies displayed a broad outline of the duration of

total caspase activation and leveling, and these results provided the period of observation for use

in the microfluidic device trials. The elucidation of the capabilities of PPIs to induce caspase-

dependent apoptosis could outline less systematically toxic therapeutic alternatives to traditional

20

cancer treatments. Furthermore, the identification of the temporal dynamics of specific caspases

activated by PPIs could prove beneficial for targeted caspase therapies, which may employ PPIs

as standalone anticancer agents.

2. MATERIALS AND METHODS

2.1 Microfluidic Device Fabrication

The procedure for the fabrication of the microfluidic devices used in this study was

adapted from the original study by Reif et al. in 2009. Poly(dimethylsiloxane) pre-polymer

(Ellesworth Adhesives) and Dow Sylgard 184 cross-linking agent (Ellesworth Adhesives) were

combined in a 10:1 ratio (w/w). The solution was mixed vigorously for five minutes, degassed in

a vacuum chamber until no air bubbles remained, and poured into a machined mold. This mold

was cured for two hours at 80°C and then cooled to room temperature. The cured PDMS was cut

into pieces approximately two inches by two inches, and adhered to a glass microscope slide

(Fisher Scientific) with additional uncured PDMS polymer. After adhering, the mold was cured

at 80°C for fifteen minutes and allowed to cool. A one-inch section of 220 µm glass capillary

(Trajan Scientific and Medical) was inserted into the channel and additional uncured PDMS was

administered to the area around the capillary and along the edges of the device to prevent

leakage. After a final curing period of fifteen minutes at 80°C, the devices were tested to ensure

proper function and stored for coating.

2.2 Coating of Microfluidic Devices with Antibody Ladder

Coating of the affinity surface was performed according to the Kuida et al. procedure,

with NeutrAvidin taking the place of streptavidin to reduce non-specific binding (Reif et al.,

21

2010). Tubes were fashioned from 30-gauge Teflon tubing (SGE Analytical Science) and 200

series barbs (Cole Parmer Instrument) for the administration of liquid. LEUR-LOK tip, 3 mm

syringes (Dickinson and Company) were utilized to push liquid into the tubing for application in

the microfluidic device. The microfluidic channel was first rinsed with 70% ethanol and 1x T50

buffer (10 mM Tris-HCl, pH 8, 50 mM NaCl). Biotinylated BSA (1mg/mL in T50 buffer)

(Fisher Scientific) was then added to the affinity channel and allowed to coat for forty-five

minutes. Subsequently, the protein was cleared from the channel with air and additional 1x T50

buffer. NeutrAvidin (0.2 mg/mL in T50 buffer) (Fisher Scientific) was then added to the channel

and allowed to coat for twenty minutes. The channel was then cleared with additional T50 buffer

and dried with air, at which point the device could be stored at 4°C for up to two months before

use. On the day of the experiment, biotinylated-anti-CD71 (50 µg/mL in PBS, pH 7.2, with 0.1%

gelatin) (Fisher Scientific) was applied to the microfluidic channel and allowed to coat for

twenty minutes. Devices were utilized immediately following the antibody coat.

2.3 Preparation of PPIs, Doxorubicin, and Fluorogenic Probe

All PPIs and the positive control drug, doxorubicin, were purchased from Sigma Aldrich.

The protocol for PPI preparation was adapted from Scaringi et al. to account for the differing

solubility of each drug. Omeprazole was dissolved in polyethylene glycol (PEG) 200 (Sigma

Aldrich) to make a 20 mM stock solution. Dexlansoprazole was dissolved in DMSO to make a

50mM stock solution. Esomeprazole was dissolved in DMSO to make a 650 mM stock solution.

The positive control drug, doxorubicin, was dissolved in DMSO to make a 20 mM stock

solution. These concentrated solutions were stored in small aliquots at -20°C and thawed just

before an experiment. All PPIs were diluted down to 1.0mM upon incubation with the cells and

22

fluorogenic probe at a total volume of 2000 µL. The fluorogenic probe, D2R (20 µM) (Sigma

Aldrich), was suspended in DMSO.

2.4 Cell Culture

Jurkat T lymphocytes were cultured in RPMI 1640 medium (Hyclone) containing 10%

fetal bovine serum (FBS) and 1% penicillin-streptomycin (Fisher Scientific) and maintained in

an incubator at 37°C and 5% CO2 atmosphere. Sub-cultures were performed every three days in

a laminar flow tissue culture hood (BSL-2) to ensure cell viability and to keep up with the

schedule of experiments. To prepare for experimentation, cells were centrifuged twice at 4000

rpm for five minutes in an Eppendorf microcentrifuge 5425, with a resuspension in 1x

phosphate-buffered saline (PBS) occurring in-between centrifugations. After washes, the cells

were transferred to a 35 mm petri dish (Greiner) for incubation with the target drug.

2.5 Baseline Caspase Activation Study

A baseline study was conducted to confirm that the selected drugs induced caspase

activity in any capacity. Experimental samples were run synchronously with a positive and

negative control, utilizing the same Jurkat cells for each sample. Fifteen milliliters of Jurkat cells

were harvested, washed twice with 1x PBS, re-suspended in three milliliters of complete

medium, and divided amongst three petri dishes. The selected PPI drug was added to the

experimental petri dish at a concentration of 1.0 mM. Doxorubicin was added to the positive

control dish at a concentration of 100 µM. No PPIs were added to the negative control dish. For

all samples, 6 µL of D2R was added directly to the culture dish (2 µL/mL). The probe was added

at the same time as the PPI drug for the experimental dishes. The six-hour incubation period

23

began immediately following the addition of the fluorogenic probe to each sample. Equivalent

amounts of either PEG 200 or DMSO were added to the negative control dishes to be congruent

with the amount of drug solvent in the experimental sample. The total volume during incubation

was initially two milliliters. At time zero of this period, 200 µL of cells were collected from the

petri dishes and washed twice with 1x PBS for imaging.

During the incubation period, the petri dishes were stored in the incubator at 37°C and

5% CO2 atmosphere to maintain cell viability. At hour six, the total volume of cells was

harvested from each petri dish. These harvests were washed three times with 1x PBS, and

another 200 µL of washed cells was collected from each for imaging. The remaining cells were

re-suspended in complete medium with 6 µL of D2R at a total volume of five milliliters per dish

and placed back in the incubator. Cells were harvested from each sample in aliquots of 500 µL

for imaging at hours twelve, eighteen, twenty-one, twenty-four, twenty-seven, and thirty from

the time of exposure to the drug. All images in this study were taken with a Nikon Monochrome

Microscope Camera (DS-Ri2) connected to an inverted microscope (Eclipse Ts2R-FL). White

light images (exposure 20 ms, DIA filter, phase contrast 2) and green light fluorescence images

(exposure 1000 ms, green light filter, phase contrast 2, 100-W Hg lamp) were taken of each

aliquot. The 10x microscope objective was utilized for the baseline study to capture a greater

number of cells. These images were saved for later analysis. Baseline activation studies were

repeated in triplicate for each drug and the controls.

2.6 Temporal Detection of Caspase Activity

For the temporal dynamics trials, microfluidic devices were utilized for the capture and

observation of the cells. Fifteen milliliters of Jurkat cells were harvested, washed twice with 1x

24

PBS, and re-suspended in a petri dish in two milliliters of complete medium with the selected

PPI drug and the fluorogenic probe. The petri dish was stored in the incubator at 37°C and 5%

CO2 for the six-hour drug incubation period. Upon the completion of this period, the total cell

volume was harvested, washed three times with 1x PBS, and re-suspended in complete medium

containing the fluorogenic probe. The cells remained in the incubator for an additional fifteen to

eighteen hours before being harvested for device observation. After the drug incubation period, a

microfluidic device was coated with anti-CD71 for twenty minutes. This device was connected

to a Y-junction prepared from Teflon tubing, and secured to the microscope stage. The

experimental cells were harvested, washed twice with 1x PBS, re-suspended at a final volume of

one milliliter, and transferred to a syringe. During the antibody coating period and cell

centrifugations, a solution of FBS-PBS was prepared. One milliliter of 10x FBS was combined

with one milliliter of 10x PBS. This solution was diluted to ten milliliters, at which point three

milliliters were collected, combined with 6 µL fluorogenic probe, and transferred to a syringe. A

third syringe of 1x PBS was also prepared. Once the cells were introduced to the channel, the

cell syringe was swapped for the 1x PBS syringe. The FBS-PBS and 1x PBS syringes remained

attached to the pumps for the duration of the experiment, and alternated pushing liquid through

the microfluidic channel. The 1x FBS – 1x PBS syringe was needed to maintain cell viability and

expose the cells to the probe. The 1x PBS syringe was required to clear excess fluorogenic probe

from the observable cell field to reduce background fluorescence during analysis.

After device coating and cell washing, the syringes were connected to the Y-junction and

mounted to automated syringe pumps (Kd Scientific). The experiment was limited by the

presence of only two pumps, so the cell syringe was attached for long enough to flood the

channel with a sufficient quantity of cells, and then it was swapped for the PBS syringe. When at

25

least one hundred observable cells were present in the channel and the syringes were secured, the

cells were allowed a twenty-minute binding period. This binding period was essential for the

adherence of cells to the affinity surface. After the binding period, the FBS-PBS pump was run at

1.5 mL/hour for three minutes to remove unbound cells from the device. This also flooded the

channel with the solution containing the fluorogenic probe. Since an excess of fluorogenic probe

solution would cause too much background fluorescence for imaging, the PBS syringe was then

run at 0.25 mL/hour for ten minutes. Following the PBS wash, the time zero imaging was

conducted, marking the beginning of the six-hour observation period. During this period, images

were taken every twenty minutes. Between images, the FBS-PBS syringe was first run for ten

minutes at 0.15 mL/hour, followed by the PBS syringe for ten minutes at 0.25 mL/hour. This

time course for the pumps ensured cell viability and sufficient exposure to the fluorogenic probe

while also maintaining the low fluorescence intensity of the background. The white light and

green light images were saved for later analysis.

2.7 Binding Affinity Study

To determine if PPI treatment affected the ability of cells to bind to the affinity surface, a

binding study was conducted. Cells were harvested and incubated using the incubation protocol

from the baseline activation study, including the addition of the fluorogenic probe and the re-

suspension in complete medium after the incubation period to remove the PPIs from the

suspension. A 500 µL aliquot of cells was removed and washed with 1x PBS every six hours for

imaging on a microscope slide and on a microfluidic device. A single set of white light and green

light images was taken for each sample on the microscope slide to verify the presence of

fluorescence. The remaining sample cells were introduced to the microfluidic device channel and

26

allowed to bind for a period of twenty minutes, at which point pre-wash white light and green

light images were taken. The channel was then washed with 1x PBS for three minutes to remove

unbound cells. Post-wash white light and green light images were then taken. Each experimental

sample was accompanied by a negative control with no added PPI, to be examined congruently.

The experimental sample and the conjugate negative control sample utilized the same cells. The

same microfluidic device was utilized for all three experiments, which were run back-to-back.

Prior studies verified the reusability of the microfluidic devices over a period of three days with a

re-coating of anti-CD71 in-between trials. To establish the percentage of cells that successfully

bound to the channel, the post-wash cell count was compared to the pre-wash cell count taken

from the respective white light images. The presence of fluorescence in the pre-wash images

versus that observed in the post-wash images was also noted to determine if cells exhibiting PPI-

induced caspase activity were selectively unable to bind to the channel.

2.8 Analysis of Images

All white light and green light images were processed utilizing ImageJ software (v. 1.4.1,

National Institutes of Health). To establish the threshold of background of fluorescence, sections

of the images that contained no cells were measured for the mean intensity of fluorescence. The

white light images provided the guidelines for selecting areas with no cells present. Several

sections of the background were measured and averaged to perform a background subtraction.

The fluorescence intensity of each individual cell in every image was measured and the

background subtraction was performed to determine if the cell was apoptotic. To create an even

more robust threshold for apoptotic activity, the fluorescence intensities of cells determined to be

non-fluorescent using the background subtraction method were utilized to create a secondary

27

apoptotic threshold. This calculation, which was based on the limit of detection equation, took

the mean plus three times the standard deviation to establish a more accurate threshold. Cells

which surpassed this threshold were considered apoptotic, showing general caspase activation.

For the baseline activation studies, every image was processed individually, with its own

unique average background. The temporal trial images were converted to a stack of images in

ImageJ, and the plugin MeasureStack was utilized to measure every image in the stack to

monitor the fluorescence of individual cells over the course of the experiment. This allowed for

the determination of the onset time and duration of caspase activity. White light images taken

from the binding affinity study were analyzed by counting pre-wash and post-wash cells utilizing

the multi-select tool in ImageJ. For the green light images from the binding affinity study,

individual apoptotic thresholds were established for the pre-wash and post-wash images to allow

for a comparison of the quantities of caspase-exhibiting cells that were observed.

2.9 Safety Considerations

Jurkat cells are classified as biosafety level one (BSL-1) organisms. Proper guidelines for

handling and sanitation and precautions for blood borne pathogens were followed. Sub-culturing

and harvesting of the cells was conducted in a laminar-flow hood, BSL-2. Gloves, lab coats, and

eye protection were worn when applicable. Any item that encountered cells was cleaned with

70% ethanol prior to disposal in a biohazard waste bin. Items that were re-used for multiple

studies, including syringes utilized for PBS, and the glass micro-capillary utilized in the devices,

were cleaned thoroughly with ethanol prior to re-use. Ethanol, which is highly flammable, was

kept away from sources of heat. Proper handling of DMSO was conducted when applicable to

prevent the absorption of harmful chemicals into the skin. Syringes lacked needles, eliminating

28

the need for sharps disposal. Care was taken to ensure that the LED fluorescent lights in the

microscope did not pose an optical threat. The positive control in these studies, doxorubicin, is

an antineoplastic chemotherapy drug that is mutagenic, carcinogenic, and teratogenic. Since the

doxorubicin was also housed in DMSO, extra care was taken to prevent exposure and absorption

through the skin. Nitrile gloves and proper protective clothing and goggles were worn when

handling doxorubicin. When possible, doxorubicin was handled in a biosafety cabinet to limit the

possibility of inhalation. The entire handling area was wiped down with soapy water following

use.

3. RESULTS

3.1 Baseline Caspase Study

To determine the general window of caspase activity following the six-hour

incubation period, a baseline caspase study was conducted for each drug. The results of the

baseline studies were utilized to select a six-hour window for the device temporal dynamics

trials. To measure when caspase activity began following treatment, the percentage of apoptotic

cells in each fluorescence image was determined utilizing the background subtraction and non-

fluorescent cell thresholds. The percentages of apoptotic cells in each image for the three drugs,

as well as the positive and negative controls, are displayed in Figure 9. Sample images for each

drug and the controls are shown in Figure 10. Aggregation was observed in each trial of

dexlansoprazole and esomeprazole, but not seen with omeprazole or the positive control. All

three drugs showed increased caspase activity over time, which was extrapolated from the

percentage of apoptotic cells observed at each timestamp. For dexlansoprazole, general caspase

activity was shown to reach 50% apoptotic cells between hours eighteen and twenty-one; this

29

corresponded to twelve and fifteen hours after the incubation period.

Dexlansoprazole also showed comparative caspase activity to cells treated with the

positive control, doxorubicin (100 µM), at hour 30 (Figure 9). Cells treated with omeprazole

reached 50% apoptosis between hours twenty-one and twenty-four, corresponding to fifteen and

eighteen hours post-incubation. Esomeprazole showed heightened activity between hours

twenty-four and twenty-seven, or eighteen and twenty-one hours post-incubation. The positive

control, cells treated with doxorubicin, showed very high caspase activity within the first six

hours of the experiment (Figure 9), which persisted through hour thirty. Cells in the negative

control group showed markedly lower caspase activity until hour thirty of the experiment. These

initial results were utilized to obtain the window of activation, the six-hour period preceding

when the percentage of apoptotic cells reached fifty-percent, for each drug.

Figure 9: Percentage of apoptotic cells by hour for each drug. Error bars represent standard

deviation for three trials. Percent apoptosis corresponds to general caspase activity. Experimental drugs show increased caspase activity over time, with dexlansoprazole showing comparable

activity to the positive control drug, doxorubicin (100 µM), at hours 27 and 30.

30

Figure 10: T=0 fluorescence images (a-e), T=30 fluorescence images (f-j), and T=30 white light images (k-o) for each sample. All experimental drugs showed visible fluorescence by hour thirty,

compared to the negative control. Dexlansoprazole and esomeprazole showed a high degree of aggregation that was not observed with the other samples. Scale bar indicates 0.25 mm.

Fluorescence images enhanced for clarity.

3.2 Microfluidic Device Temporal Trials

Appropriate six-hour windows where caspase activation might occur were selected for

each drug. For dexlansoprazole, cells were observed in a microfluidic device from hours fourteen

to twenty-two. Omeprazole-treated cells were observed from hours eighteen to twenty-four.

Cells treated with esomeprazole were observed from hours twenty-two to twenty-six. The results

of these selected windows showed no evidence of caspase activation after repeated trials. It was

theorized that, due to the microenvironment being much different in the device than in the

culture dish, caspase induction and subsequent apoptosis were occurring either much earlier or

much later than the windows viewed in the baseline study.

To check for caspase activity within the thirty-hour window utilized in the baseline study,

cells treated with each drug were observed on devices from zero to six, six to twelve, twelve to

31

eighteen, eighteen to twenty-four, and twenty-four to thirty hours. For every six-hour time-

period, all three experimental samples showed no detectable fluorescence (Figure 11). This was

compared to the positive control, doxorubicin-treated cells, which had a caspase onset time of

3.66 ± 1.67 hours (from three experiments, with n = 100 cells), and the negative control, which

showed no significant fluorescence in any time window. Additionally, microfluidic device

temporal trials displayed a low percentage of experimental cells that successfully bound to the

affinity surface. It was hypothesized that PPI exposure alters the cellular surface receptors in

some capacity, reducing the ability of the treated cells to bind to the device’s channel. A binding

study was necessary to confirm that this phenomenon was specific to PPI-treated cells.

Figure 11: Fluorescence images and accompanying end-stage white light images from the microfluidic device temporal trials. Timestamps are provided for each image. Doxorubicin

shows a high degree of fluorescence by hour six of the experiment, while the experimental drugs show very few fluorescent cells throughout thirty hours of experiments. Experimental drug

images are from different trials, each spanning six hours for a total of thirty hours of observation per drug. Images enhanced for clarity.

32

3.3 Binding Affinity Study

The results of the binding study confirm the correlation between PPI-exposure and lower

binding affinity. Binding percentages calculated from the respective pre-wash and post-wash

images at each time interval are shown in Figure 12, with sample pre-wash and post-wash

images displayed in Figure 13. Cells treated with any of the three PPI drugs showed a

measurably lower binding affinity that tended to decrease over time. Notably, experimental drugs

showed binding affinities comparable to the negative control at hour zero upon initial exposure

to the PPI drugs. However, by hour six of incubation with the drugs, experimental cells showed

approximately a 50% decrease in binding affinity. The negative control was run in triplicate,

with each trial accompanying one of the experimental samples, and the results of the three trials

were averaged. The control was shown to maintain roughly the same binding affinity throughout

the thirty-hour experiment, as was consistent with anticipated results. To further eliminate

possible sources of discrepancy between the binding affinities of the experimental samples and

the controls, the same microfluidic device was utilized for every experiment. A previous binding

study had shown the antibody coat to remain viable over the course of a week if the device was

washed with 1x PBS in between experiments; to ensure the function of the antibody coat, these

experiments were run consecutively over the course of three days. Additional images were taken

showing the fluorescence of the cells pre-wash and post-wash (Figure 14). These images were

utilized to assess whether cells that were unable to remain bound were also expressing caspase

activity.

33

Figure 12: Percentage of bound cells for each drug by hour, taken from the percentage of cells leftover after the PBS wash. PPI-treated cells all show a dramatically reduced binding affinity

immediately following the 6-hour drug incubation period. This phenomenon is not observed with the negative control, the data for which was averaged from three separate trials. Each individual negative control accompanies one of the experimental groups. The same cells were used for both the experimental sample and its respective negative control. All samples were tested on the same

microfluidic device, providing negative control data in triplicate. Error bars represent the standard deviation from the three negative control trials; experimental drug trials were conducted

once each.

34

Figure 13: Sample pre-wash and post-wash images for each drug and the negative control. These images were taken twenty-four hours into the experiment for each sample. Cells are

numbered in green. The negative control shows roughly the same quantity of cells in both the pre-wash and post-wash images, whereas the experimental samples show a significant reduction

in cell number. This translates to a binding affinity that remains unchanged for the negative control, but is lowered by approximately 50% in the experimental samples. Scale bar represents

0.25 mm.

Figure 14: Pre-wash and post-wash fluorescence images for each drug, corresponding to the white light images in Figure 13. The negative control was not utilized as a comparison here

because there was no fluorescence observed. Experimental drugs appear to lose roughly half of the fluorescent cells that are present prior to the PBS wash. It is currently unclear whether lack of

binding is correlated to caspase activity, or whether it is a random effect. Arrows indicate residual PDMS that is exhibiting fluorescence. Images enhanced for clarity.

35

4. DISCUSSION

The results of the baseline caspase study revealed the approximate window where

caspase activity increased for each drug, allowing for the selection of the six-hour period for the

temporal trials. Given that this study served as a preliminary assessment, it was performed in

triplicate to verify the results of each trial. The caspase study confirmed that all three of the PPI

drugs are capable of inducing caspase-dependent apoptosis following six hours of exposure to

1.0 mM of drug (Figure 9). The 1.0 mM concentration for the PPIs was selected from a prior

study by Scaringi et al., which showed measurable levels of apoptosis in Jurkat cells following

exposure to this concentration. Dexlansoprazole showed the most promising caspase activity out

of the three experimental drugs; it displayed comparable activity to the positive control,

doxorubicin, at hours twenty-seven and thirty. While the concentration of the dexlansoprazole

was much higher than that of doxorubicin (1.0 mM versus 100 µM), the achievement of a

comparable level of activity is noteworthy.

Doxorubicin is one of the most powerful chemotherapeutic drugs ever administered in a

clinical setting, and can kill cancer cells at any stage of maturity (Johnson-Arbor and Dubey,

2021). It has been utilized in clinical practice since the 1960s for the treatment of acute

lymphoblastic leukemia, acute myeloblastic leukemia, Hodgkin lymphoma, and small cell lung

cancer. Doxorubicin’s primary method of action is intercalation within the DNA, inhibiting DNA

and RNA synthesis. It also inhibits topoisomerase II, leading to DNA damage that induces

mitochondrial apoptosis, alongside free radical-mediated cell death and ferroptosis (Johnson-

Arbor and Dubey, 2021). Its mechanism of inducing cellular death via mitochondrial-mediated

apoptosis and its non-cell-cycle-specific targeting makes it a comparable positive control to the

PPI drugs. Despite the concentration difference between the positive control and the

36

experimental PPIs, the incidence of dexlansoprazole showing comparative activity to that of

doxorubicin confirms its potential for further evaluation as a standalone therapeutic.

Since D2R is utilized to monitor general caspase activity, the window of activation for the

baseline study encompassed both the initiation and execution stages of apoptosis, rather than

highlighting the activity window for specific caspases. The D2R probe is suitable for the

detection of both the intrinsic and extrinsic pathways of apoptosis (Martinez, 2010). Care was

taken in selecting the appropriate six-hour observation period for temporal trials from the

baseline study, since initial caspase activity is present even in the negative control group.

Apoptosis is a normal process that occurs constantly in a healthy population of cells (Elmore,

2007), meaning that some level of caspase activity should be observed for any given sample of

cells. All three of the PPI drugs showed less than twenty percent apoptotic cells in the first

twelve hours of the experiment.

The higher levels of activity seen in the negative control at hour thirty are likely

attributable to low cell and medium volume in the culture dish by the end of the experiment.

Once the cells were re-suspended in complete medium after the incubation period, no additional

media changes were performed so as not to disturb the cells. The removal of aliquots of cells for

imaging reduced the total volume leftover in the dish, potentially causing the remaining cells to

use up available nutrients in the medium and begin to undergo starvation. These cells may also

be undergoing apoptosis as a result and would be included in the measured levels of caspase

activity. However, the experimental samples for each drug show elevated caspase activity much

earlier on in the timeline and with a notably higher intensity than the negative control.

Furthermore, the levels of apoptosis seen in this study differ from normal levels of apoptosis that

are observed in a healthy population of Jurkat cells. Jurkat cells are an immortalized line of

37

lymphocytes that were derived from a 14-year-old boy with T cell acute lymphoblastic leukemia

(Bartelt et al., 2009). Immortalized cell lines can proliferate indefinitely, so a very small number

of cells will actively undergo apoptosis in a culture of Jurkat cells provided they are given

sufficient medium and growth conditions (Bartelt et al., 2009). Two-tailed, paired t-tests

performed with the experimental drug versus the negative control at each time point showed that

the effect of each drug on apoptosis levels was statistically significant (p < 0.0001 for n = 100

cells, with a confidence interval of 95%). This finding indicates that most of the apoptotic

activity seen in the experimental cells is the result of PPI exposure triggering caspase activation.

These results revealed that caspase-dependent apoptosis in Jurkat cells, resulting from

PPI treatment, is a late-onset process that does not intensify for at least twelve to fifteen hours

following the incubation period. This finding contrasts the early onset time seen in the positive

control, doxorubicin, which showed approximately 90% caspase activity within the first six

hours of exposure (Figure 9). Doxorubicin has an elimination half-life of at least twenty and up

to forty-eight hours (Gabizon et al., 2003 and Johnson-Arbor and Dubey, 2021). Omeprazole has

an elimination half-life of thirty minutes to one hour (Cederberg et al., 1989). Esomeprazole has

a half-life of one to one and a half hours. Dexlansoprazole has a dual delayed-release formula

intended to create two peaks of therapeutic activity, and a half-life of one to two hours (Strand et

al., 2017).

Dexlansoprazole is designed to become active first at a pH of 5.5, and later at a pH of

6.75 (Gasiorowska, 2017). The cancer cell microenvironment pH ranges from 5.6 to 6.8 (Lin et

al., 2019). Omeprazole and esomeprazole become active just once at a pH of approximately 5.4

(Navarro et al., 1998 and Kim et al., 2019), meaning that the average cancer cell’s

microenvironment may not be acidic enough to trigger the activation of these drugs. Since

38

dexlansoprazole’s secondary activation is designed to take place at 6.75, it is much more likely

to be triggered by the average cancer cell. Given that dexlansoprazole showed the greatest

efficacy of inducing caspase activity out of the experimental drugs, it may be hypothesized that

its dual-release formula, with the second release occurring at a higher pH, enhances its ability to

trigger apoptosis in Jurkat cells. This is further evidenced by dexlansoprazole’s comparative

caspase activity to that of doxorubicin at hour thirty (Figure 9).

This study conducted an acute drug exposure rather than a chronic, multi-day exposure.

Clinical administration of doxorubicin is normally performed intravenously in twenty-one day

intervals (Johnson-Arbor and Dubey, 2013), meaning chemotherapy requires chronic exposure to

the drug. Doxorubicin still showed effectiveness in this acute exposure due to its capabilities as a

potent antineoplastic agent. The concentration of each PPI drug was maintained at 1.0 mM for all

treatments, and direct incubation of the cells with the target drug occurred for six hours. This

concentration is very high, and likely much higher than what would normally be prescribed in a

clinical setting. The goals of this study were to assess the apoptotic induction capabilities of the

PPIs in an acute exposure, and future experiments may be conducted to assess whether the PPIs

can induce caspase activity in a chronic exposure over multiple days or weeks at a lower

concentration. Since chemotherapy is normally carried out for several months or indefinitely to

eliminate and control tumor growth, these PPIs could be tested in a similar schedule as common

chemotherapeutics to determine their effectiveness. Dexlansoprazole may be particularly

effective in a chronic exposure due to its dual-release mechanism, meaning it is able to maintain

therapeutic levels in the body for an extended period compared to the other PPIs. Determining

the temporal dynamics of caspase activation on an individual-cell basis may enhance the ability

to outline a successful chronic exposure, highlighting the need for viable microfluidic device

39

temporal trials in future studies.

The initial windows of activation selected from the baseline study showed very little

fluorescence observed for each sample of treated cells. Doxorubicin showed a high degree of

caspase activity within the first six hours of exposure (Figure 11), confirming that the structure

of the device allows for the occurrence and visualization of fluorescence. It is theorized that the

baseline study timeline for activation differs from that of the device due to the difference in

conditions of the micro-environment to a culture dish. The microfluidic device represents an

environment that is more physiological in nature, as it allows for gas exchange and provides a

constant allotment of moving medium. Cells bound to the affinity surface can expunge toxins

and waste that is carried away by the movement of the medium; cells in the culture dish must

remain suspended in their own waste. It is also possible that the culture dish cells show greater

accumulation of D2R over time as the cells are in static medium, and diffusion of the cleaved,

fluorescent R110 molecule out of the cell may have occurred more slowly. This may have led to

an amplified fluorescence signal long after the caspase cascade had already begun.

Troubleshooting the caspase activation window for the experimental drugs required five

separate temporal assays each, to search the entire window of activity observed in the baseline

study. These assays monitored cells during the incubation period and up to twenty-four hours

following the incubation period. To directly observe cells during the six-hour incubation period

with the target drug, the PPI was added to the FBS-PBS-D2R and PBS syringes at a

concentration of 1.0 mM. The drug was added to both syringes to ensure the cells were

constantly exposed to the drug throughout the course of the experiment. Cells were exposed to

the fluorescent probe consistently for the duration of the experiment, apart from the five-minute

periods of PBS washing that preceded every imaging round. Despite constant drug and

40

fluorescent probe exposure, none of the selected time windows showed significant caspase

activity. Several factors may influence the absence of fluorescence seen in the microfluidic

device temporal trials, with pH being the common denominator. PPIs are inactive in basic

conditions, but become active as inhibitors in an acidic environment (Walsh, 2015). In the

baseline activation studies where cells were suspended in medium in a culture dish, the waste

products excreted by the cancer cells accumulated in solution, resulting in the acidification of the

extracellular environment.

The PPI drug added to this dish would become active in this environment, conferring its

caspase induction ability. However, in the microfluidic device temporal trials, the cells are

subjected to exposure to a constant influx of PBS at a pH of 7.4, meaning the channel

environment was slightly alkaline throughout the duration of the experiment. This may have

counteracted the microenvironment acidification of the cancer cells, resulting in the inactivation

of the PPI drugs. This would mean the drugs’ apoptotic induction capabilities were not taking

effect, and would result in an absence of caspase activity. In the future, an assessment of the pH

of the culture dish over the course of the thirty-hour experiment will be desirable for confirming

this link. If the culture dish is shown to have a drastically lower pH than the 7.4 PBS, additional

microfluidic device trials should be conducted with an acidic PBS buffer to determine if caspase

activity is present. If so, this would mean that PPIs may only trigger the induction of the caspases

if the extracellular environment is maintained at an acidic pH.

Cells treated with PPIs showed a lower binding affinity to the microfluidic device’s

antibody ladder than the negative control (Figure 12). In current applications, PPIs target the

H+/K+-ATPase system, a specific V-ATPase, found at the secretory surface of gastric parietal

cells (Masaoka, 2008). It is known that V-ATPases play a role in the regulation of tumor pH, and

41

V-ATPase is overexpressed on the cell surface in multiple cancer cell lines (Chueca et al., 2016).

Since PPIs target proton pumps located on the cellular surface, it is not unreasonable to expect

the cell membrane to be affected in other ways by exposure to these drugs. The antibody utilized

in this study, anti-CD71, is utilized for the capture of cells expressing the transferrin receptor

(TfR). TfR, a 95 kDa homodimeric type II transmembrane glycoprotein, regulates cell growth

(Loisel et al., 2011). TfR is overexpressed in multiple cancer cell lines (Kasibhatla et al., 2005),

meaning this antibody is a useful tool for cell affinity chromatography studies, and shows a high

binding affinity for multiple types of cancer. Anti-CD71 is not known to induce apoptosis in

Jurkat cells, making it ideal for studies where cell capture for the purposes of inducing apoptosis

via another drug is desired (Reif, 2009).

Although TfR expression is known to be growth-dependent, its expression in Jurkat cells

can be predicted based on the sub-culturing schedule. TfR is expressed at low levels in non-

proliferating cells, but is intensely upregulated during proliferation (Brell et al., 2020). Since

Jurkat cells have a doubling cycle of approximately one-to-two days (Schoene and Kamara,

1999), cells were selected for the microfluidic device trials three-to-four days after sub-culturing

to maximize expression of TfR. This means that the lowered binding affinity of the experimental

cells is not attributable to low expression of TfR alone, especially with the negative control of

the same cell population exhibiting normal levels of binding. An alternate antibody, anti-TCR,

was utilized in the initial temporal dynamics trials, but was shown to have a poor binding affinity

for the cells after the six-hour incubation with the target drug, even when cells were harvested at

various initial growth stages. Anti-TCR binds cells expressing the T-cell receptor protein on the

cellular surface and has been utilized widely in conjunction with Jurkat cells (Abraham, 2004).

Unlike TfR, the T-cell receptor protein is not growth-dependent, and should not suffer from the

42

same obstacles in binding that may accompany the anti-CD71 antibody. Since both proteins are

expressed on the outer membrane of the cellular surface, and both antibodies displayed lower

binding percentages in the experimental samples, it can be theorized that the TfR and T-cell

receptor proteins are affected in some way by PPI exposure, reducing their density on the cellular

surface and resulting in a lower binding percentage for PPI-treated cells. Exposure to the PPIs

may also alter the physical shape of the membrane, potentially preventing the proteins from

being exposed to and contacting the antibody ladder on the microfluidic device.

The aggregation observed in cells treated with dexlansoprazole and esomeprazole (Figure

12) was consistent throughout the repeated trials, and most often began after hour twelve of the

experiment. This aggregation may play a role in the reduction in binding affinity seen in the cells

following PPI exposure. Aggregation was not present in the experimental cells prior to treatment

with the PPIs, and was not found in the doxorubicin-treated cells or the negative control. This

suggests that dexlansoprazole and esomeprazole induce this effect directly, although the

mechanism for this action is unknown. Aggregation does not appear to be correlated to the

inhibitor drug’s ability to induce apoptosis. While dexlansoprazole showed a high degree of

aggregation and was also the most potent drug, esomeprazole showed comparative degrees of

aggregation, but was the least potent of the three drugs. Omeprazole, which showed no evidence

of aggregation in any of the studies, induced higher levels of apoptosis than esomeprazole.

An important distinction between the aggregated and non-aggregated cell groups was the

solvent utilized to dissolve the PPIs. Dexlansoprazole and esomeprazole were both dissolved in

DMSO, while omeprazole was dissolved in PEG 200. The selection of the PEG 200 solvent for

omeprazole was made according to the study by Scaringi et al., and was intended to replicate the

methods outlined in that study. However, dexlansoprazole and esomeprazole are highly insoluble

43

in PEG 200, and were instead dissolved in DMSO. Since doxorubicin was also dissolved in

DMSO and showed no aggregation, there is no evidence to suggest that the DMSO was

contaminated. However, this does indicate a possible link between PPI exposure and aggregation

that may be disrupted by the presence of PEG 200. While PPIs are not known to directly induce

cell aggregation, they are known to enhance platelet aggregation and clot formation (Wu et al.,

2019) in patients with bleeding peptic ulcers.

Their ability to enhance aggregation of the platelets may also be present in cancer cells,

resulting in the treated cells sticking to each other in a manner not seen in untreated cells. This

phenomenon remains unconfirmed and largely unexplored. However, PEG 200’s role in

eliminating this aggregation in the omeprazole-treated cells may be speculated. PEG derivatives

are known to be capable of modulating protein aggregation (Rajan et al., 2006). If the PPIs are

altering the cancer cells’ surface receptor proteins and leading to aggregation, the PEG 200

solvent the omeprazole is housed in may modulate this ability, leading to the absence of

aggregation. In future studies, cells may be treated with omeprazole dissolved in DMSO to

determine if aggregation occurs. The aggregation may be responsible for the low binding

affinities observed in the binding study. If the link between aggregation and PPI exposure can be

solidified, methods may be produced that can circumvent the reduction in binding affinity,

allowing for the completion of successful microfluidic device temporal dynamics trials.

The use of the microfluidic devices for individual-cell analysis has many benefits.

Primarily, the ability to observe the caspase activity onset and duration of a single cell allows for

a much more precise characterization of the temporal dynamics. However, the initial lack of

fluorescence and the low binding affinity of the experimental samples presented challenges to

the viability of these experiments. It was hypothesized that the window of caspase activation

44

could be somewhere within the thirty-hour time-frame, but cells expressing caspase activity were

selectively unable to bind. Without sufficient binding, and with the possibility that cells

exhibiting caspase activity were selectively unable to bind, cells could not be observed for

temporal analysis utilizing the microfluidic devices. These results warranted a binding affinity

study to assess whether the link between PPI exposure and lower binding percentages was

significant. If this link could be validated, it could allow for the selection of an antibody whose

antigen remains unaffected by PPI exposure.

The results of the binding study strongly suggested a link between PPI exposure and

reduced binding affinity of the cells to the affinity surface (Figure 12). All three of the

experimental drugs showed a significant decrease in the percentage of cells that successfully

bound to the device following the six-hour incubation period. The experimental cells showed a

binding affinity roughly the same as that of the negative control at hour zero, prior to the

incubation period. The negative control cells maintained their binding affinity at roughly eighty-

percent throughout the thirty-hour experiment. To ensure that a comparison was possible, the

same cells were utilized for the experimental samples and their respective controls. The same

microfluidic device was utilized for all three of the consecutive experiments, with a re-coating of

anti-CD71 occurring every twenty-four hours to maintain viability. These results suggest that the

lowering in observed binding affinities is attributable to PPI treatment. Since this experiment was

only conducted with a singular trial for the experimental cell groups, additional replicates are

needed to verify results.

With respect to the temporal dynamics assays originally intended to be performed, the

binding affinity study presents an obstacle. With the drastic reduction in the binding affinity of

cells post-treatment, it becomes more difficult to capture a suitable quantity of cells for caspase

45

detection. The reduction in binding affinity may be related to the lack of fluorescence observed

in the microfluidic temporal trials over the course of the thirty-hour period. Fluorescent and non-

fluorescent cells are both shown to be subject to removal during the wash (Figure 14), in what

appears to be equal likelihoods, meaning that the reduction in binding affinity appears to be

random and not correlated to the presence of caspase activity in the cell.

Another possibility is that non-fluorescent-apparent cells that cannot bind may be

undergoing apoptosis, but are not yet exhibiting enough fluorescent for detection by the time

points in the binding study. If caspase-expressing cells are selectively unable to bind to the anti-

CD71 antibody ladder following exposure, this renders the use of the microfluidic devices for

single-cell observation unviable as of now. In the future, the identification of an antibody that

maintains cellular binding capabilities following PPI exposure would allow for the microfluidic

temporal trials to continue. Obtaining a suitable antibody would also allow for longer

microfluidic device experiments, allowing for the characterization of the entire duration of

caspase activity on an individual-cell basis. Identifying the duration of caspase activity seen in

both doxorubicin and the experimental drugs could be beneficial for outlining a complete

timeline for caspase induction and activity, furthering the possibilities of targeted caspase

therapies involving PPIs. The selection of a different antibody may offer additional benefits as

well.

The selection of the anti-CD71 antibody, which targets only the TfR receptor, was made

to reduce non-specific binding wherever possible, while maintaining a high binding affinity to

allow for a large quantity of observable cells. This antibody has displayed growth-dependent

variations in binding affinities in previous studies (Brell et al., 2020). To ensure that a sufficient

quantity of cells was being captured during the temporal trials, sub-cultures were performed on a

46

schedule to reduce the number of discarded cells and conserve medium. Cells were utilized three

days after each sub-culture, when the colony displayed a golden-orange color (Figure 15) and a

high confluency. The cells were shown to give the best binding affinity when in this stage of

growth. Failure to follow the strict sub-culturing schedule led to failed experiments due to a low

quantity of observable cells. Successful microfluidic device experiments were required to have at

least one hundred bound and observable cells to reduce standard error and ensure validity of

results. For these trials to produce viable results, the alternate antibody must show a high binding

affinity that is not hindered by PPI exposure.

Figure 15: Optimal cell color utilized in the microfluidic device trials, including the binding

affinity study and the initial temporal analysis trials. Sub-cultures were performed weekly, with cells incubating for at least three days following sub-culturing before they were used in any

experiments. This allowed for optimal expression of growth-dependent TfR for binding.

The microfluidic devices presented another experimental challenge. Due to the sensitivity

of the glass micro-capillary and the likelihood of the channel malfunctioning, the temporal

experiments had a success rate of approximately fifty percent. The most common source of

47

device failure was residual PDMS polymer present in the microfluidic channel (Figure 16),

which led to channel obstructions or the development of bubbles which removed bound cells.

Bubbles were also produced in the PBS and FBS-PBS syringes. When a bubble entered the

channel, the field of vision was impeded, and most bound cells were removed (Figure 17). In

cases where cells remained bound but bubbles could not be dislodged from the field of vision,

the bubbles impeded analysis and made the contrast of the fluorescence images unbalanced

(Figure 18). This presented a major problem for this study, as the six-hour timeframe was critical

to elucidating the temporal dynamics. Once cells were removed from the channel by a bubble,

the experiment had to be cancelled, resulting in wasted resources. Restarting the six-hour

temporal observation was not possible, as the cells were on a schedule that began with the PPI

incubation. Restarting the experiment would disrupt the sensitive timeframe. In future studies

when possible, the use of machined devices or the addition of a de-bubbler in the tubing setup

could increase the success rate of the temporal trials.

Figure 16: Residual PDMS at the point where the glass micro-capillary meets the opening of the microfluidic channel. The glob of PDMS impedes the flow of liquid from the capillary into the

channel, rendering the device unusable.

48

Figure 17: A section of the microfluidic device channel that was prone to the development of bubbles (top) and a later image of a bubble obstructing the entire field of vision (bottom). This phenomenon was common and unfortunately led to the failure of multiple experiments, which

had to be redone.

Figure 18: A bubble in the field of vision of an unenhanced fluorescence image. The presence of the bubble makes background subtraction impossible as it imbalances the average contrast of the

entire image.

49

Strengthening the microfluidic devices and eliminating bubbles would also allow for

extended-view trials. These trials would be beneficial for capturing the entire window of caspase

activity, provided that a suitable antibody could be selected for use in the cells exposed to the

PPIs. Doxorubicin, which had an average onset time of 3.66 ± 1.67 hours, showed persistent

caspase activity throughout the remainder of the six-hour experiment following the onset. The

standard six-hour microfluidic device experiments were too short to reliably depict the entire

window of caspase activity. A more robust microfluidic device setup would allow for an

extension of the observation period to a total of nine hours, allowing for the characterization of

the entire timeline of caspase activity following doxorubicin exposure. This would be a valuable

comparison tool for the caspase activity durations of other anti-cancer agents, including PPIs, to

further the possibilities of developing targeted caspase-therapies in the future.

This study verified that all three experimental PPI drugs induce late-stage apoptosis in

Jurkat cells via the use of the D2R general caspase probe. This study confirmed that the

fluorescence observed in the positive control and PPI-treated cells was directly proportional to

caspase activity, since the D2R probe is suitable for the detection of general caspase activity.

However, it is also worth noting that caspases have been implicated in the pathways of some

forms of programmed necrotic cell death (Yuan, 2016). Since apoptosis is desired over necrosis

due to the lack of inflammatory response and less disruption to the overall system (Kurosaka et

al., 2003), it would be useful to distinguish between the two. To further confirm that the cell

death observed in this study is due to caspase activity in apoptosis, and not necrosis or another

form of cell death, additional methods of verification are desired in future studies. Annexin V is

a protein that binds to phosphatidylserine on the plasma membrane of apoptotic or necrotic cells,

and is utilized for staining (Crowley et al., 2016). Propidium iodide is a fluorescent dye that can

50

only penetrate the cell membranes of necrotic cells. Co-staining with Annexin V and propidium

iodide would allow for the determination of which cells are apoptotic and which cells are

necrotic (Crowley et al., 2016). This procedure would verify that observed caspase activity is

solely due to apoptosis, strengthening the results of this study.

The results of this study did not attempt to examine the mechanism for how PPIs trigger

caspase activation, but this information may be desirable in future studies. In a prior study,

lysosomal damage was implicated as a factor in the onset of apoptosis following the treatment of

Jurkat cells with omeprazole (Scaringi et al., 2003). The percentage of cells with lysosomal

perturbations was shown to increase parallel to the percentage of apoptotic cells, indicating that

lysosomal damage led to the triggering of caspase-dependent apoptosis (Scaringi et al., 2003).

Since the PPIs in this study function similarly, future experiments may examine whether the

caspase activity seen in dexlansoprazole and esomeprazole is related to damage to the Jurkat

cells’ lysosomes. This examination may be performed utilizing acridine orange staining.

Acridine orange is a fluorescent dye that readily accumulates in the lysosomes; if the lysosomal

membrane is compromised, the dye will leak out into the cytosol, where it can be detected

(Scaringi et al., 2003). Determining if lysosomal damage is a key factor in triggering the caspase

cascade following PPI treatment would provide additional insights on how these drugs may act

as anti-cancer agents.

5. CONCLUSION

The purpose of this study was to verify the presence of and characterize the temporal

dynamics of caspase activity in Jurkat cells following exposure to the proton pump inhibitors

dexlansoprazole, omeprazole, and esomeprazole. Fluorescence microscopy allowed for the

51

observation of caspase activity in treated cells over a period of thirty hours on a multi-cell and

individual-cell basis. The fluorogenic probe in this study, D2R, is rapidly up-taken by the cancer

cells and readily cleaved by caspases within the apoptotic cascade. Cleavage of D2R releases the

fluorophore R110 which diffuses out of the cell very slowly. Caspase activity is thus directly

correlated to fluorescence intensity observed in the cells during real-time observation. A baseline

caspase activation study verified that all three of the experimental drugs induce significant

caspase activity in the cells following six hours of incubation with 1.0 mM of PPI drug, with

dexlansoprazole showing the highest efficacy of induction (Figure 9). Microfluidic devices were

utilized to capture cells to an affinity surface for real-time determination of caspase activation.

While the microfluidic devices were not initially suitable for the temporal dynamics studies due

to low observed fluorescence and low quantities of bound cells, these challenges led to the

development of a binding affinity study. This study showed that PPI exposure reduces the

binding affinity of the cells to the affinity surface following the incubation period (Figure 12),

indicating that PPIs alter the cell surface receptors in some fashion.

Future studies will verify that the cell death observed in this work is largely attributable

to apoptosis via Annexin V staining. Additionally, the identification of a suitable antibody for

use in the microfluidic device temporal trials will allow for a further exploration of caspase

activity on an individual-cell basis. Extended-view microfluidic device trials will allow for the

characterization of the entire duration of caspase activity resulting from doxorubicin exposure

and PPI exposure, aiding in creating a timeline for the entire caspase cascade that follows drug

treatment. This study may also be modified to lower the concentrations of the PPI drugs and

instead focus on a chronic exposure, to monitor the occurrence of caspase activity in Jurkat cells

following less-potent, but more frequent contact with the PPIs. Finally, other PPI drugs may be

52

assessed for their apoptosis-inducing capabilities, particularly those which might employ a dual-

release formula. These future studies will expand upon the results of this work, aiding in

evaluating PPIs for their potential as standalone anti-neoplastic agents.

53

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