The Impact of Alveolar Type II Cell Mitochondrial Damage and Altered Energy

Production on Acute Respiratory Distress Syndrome Development During Influenza A

Virus Infection

Dissertation

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy

in the Graduate School of The Ohio State University

Lauren May Doolittle, BA

Biomedical Sciences Graduate Program

The Ohio State University

2020

Dissertation Committee

Ian Davis, BVSc(hons), PhD, Advisor

Jacob Yount, PhD, Co-Advisor

Joshua Englert, MD

Jordi Torrelles, PhD

Copyrighted by

Lauren May Doolittle

2020

Abstract

Influenza A virus (IAV) is a respiratory pathogen that causes seasonal influenza epidemics and harbors demonstrated pandemic potential. The 1918 “Spanish Flu” pandemic was caused by a novel IAV strain and resulted in 20-50 million deaths. Today, even with access to influenza vaccines and anti-viral drugs, the World Health

Organization estimates that influenza still causes between 3 and 5 million cases of severe illness each year, resulting in 250-500,000 deaths. Many of these deaths are from acute respiratory distress syndrome (ARDS), a type of respiratory failure that develops in up to 50% of critically ill influenza patients. Although influenza is the leading viral cause of ARDS, many other illnesses, including pneumonia, sepsis, trauma, brain injury, and most recently, COVID-19, can also progress to ARDS. The all-cause mortality for ARDS is around 40%, and, regardless of initial cause, once developed ARDS can only be treated with supportive care, often involving mechanical ventilation in an intensive care unit (ICU). ARDS is associated with substantial morbidity and mortality, and there is a clear need for novel therapeutic strategies that can prevent or attenuate this condition.

To advance ARDS research, we have developed a mouse model of IAV-induced

ARDS that recapitulates the clinical signs of ARDS seen in human patients. Our work in this model focuses on the alveolar type II (ATII) cell, a pulmonary epithelial cell that is critical for maintaining lung homeostasis. Numerous ATII cell activities are altered by IAV

infection, and ATII cell dysfunction is associated with the development of ARDS in our model. We have previously shown that IAV infection results in decreased synthesis of the major cellular phospholipids phosphatidylcholine (PC) and phosphatidylethanolamine

(PE) in ATII cells, and here we investigate the impact of altered phospholipid synthesis on ATII cell mitochondrial function and ARDS development. Mitochondria are composed of two membranes, which are predominantly made up of PC and PE. We found that IAV infection changes ATII cell mitochondrial structure and membrane phospholipid composition. This results in a significant decrease in mitochondrial energy production in the form of ATP. ATII cells do not increase glycolysis to compensate for reduced mitochondrial ATP production following IAV infection, nor does reduced mitochondrial function induce apoptosis. Mitochondrial energy production can be returned to homeostatic levels by treating mice with CDP-choline following IAV infection. CDP- choline is a PC precursor, which we have shown is able to rescue IAV-induced defects in ATII cell PC synthesis. Excitingly, CDP-choline also significantly attenuates clinical signs of ARDS in IAV-infected mice. Our data suggests that this occurs through a restoration of ATII cell energy production, which facilitates ATII cell function and therefore opposes ARDS development. Overall, this work elucidates the contribution of mitochondria to ATII cell dysfunction during influenza and ARDS and proposes a therapeutic strategy to counteract these processes and attenuate disease.

Dedication

To my grandmothers, Fay Snyder Blackburn and Maria Zergenyi Doolittle.

“A wise woman wishes to be no one’s enemy;

a wise woman refuses to be anyone’s victim.”

-Maya Angelou

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Acknowledgments

Graduate research is a major intellectual, emotional and technical endeavor, and

I could not have completed my doctoral degree without help from so many amazing, brilliant, open-hearted people. Endless thanks to my PhD advisor, Dr. Ian C. Davis for 4+ years of superb mentorship, patience, and support, and for giving me a chance to be a part of a fascinating and innovative research program. You have encouraged me to master new skills, think creatively, and forge my own path in biomedical research, and I am truly grateful. Thanks for all the free and delicious tapas dinners- I expect to continue to be invited. I have also benefitted greatly from the mentorship of my dissertation committee members: Dr. Jacob Yount, Dr. Joshua Englert, and Dr. Jordi Torrelles.

Thank you all for your support and input on my research, my academic progress, and my future plans. Thank you also for your patience with my seemingly endless scheduling and logistical emails, and for letting me refer to you collectively as “The Three Js” every time I give a research talk.

To the rest of the Davis lab, I could write another dissertation about my gratitude to you all, but I will try to fit it into one paragraph. Thank you from the bottom of my heart to Dr. Lucia Rosas; I would not be the researcher I am today without your excellent technical mentorship and expertise. You are a thoughtful, skilled, meticulous and patient scientist, and I am so fortunate to have had you as a colleague throughout my PhD.

Many thanks also to my other lab mom, Lisa “The Mouse Whisperer” Joseph, another

gifted, hard-working scientist. You two have taught me everything I know about working at the bench, and we have laughed a lot along the way. I could not have done anything in this dissertation without all your help over the years. To Dr. Kate Nolan and Andrew

Nelson, thank you for choosing to spend a part of your research careers with us. You have both brought so much sass and hilarity into the lab, but also smarts, a willingness to work, and valuable experiences and insights. To top it all off, you taught me lots of animal facts, kept me up to date on vet school and OSU gossip, and I got to hang out with Tootsie. What more could a graduate student ask for? A big thank you to Adam,

Nicole, Lindsey, Hasan and Kelsey for letting me practice my mentorship skills on you; it has been so rewarding to watch you all grow as scientists and veterinarians. Thank you also to my predecessor Dr. Parker Woods for encouraging me to take a chance on the

Davis lab, and for always making time to dish out wisdom on science and on life, over a cold drink when possible. I cannot say it enough, thank you all for all your help and support over the last five years; I will miss you all when I graduate!

I feel fortunate to have even more academic role models to thank from my time at

Wake Forest. Thank you to my French professors Dr. Kendall Tarte and Dr. Sally

Barbour for being excellent educators and warm, friendly mentors, and for giving me the opportunity to study in Dijon; easily my most meaningful semester of undergrad, which, in a round-about way, led me to biomedical research. Thank you to the many Biology faculty members who opened my eyes to new areas of the field and inspired me to pursue research- Dr. Susan Fahrbach, Dr. Ray Kuhn, Dr. Pat Lord, and Dr. Brian Tague;

Wake Forest is lucky to have all of you. I am so grateful to Dr. Gloria Muday, my first

research mentor, for her unwavering commitment to high-quality undergraduate research experiences. Her mentorship prepared me to succeed at the graduate level, and I truly appreciate her patience and support in helping me reach my goal of acceptance into a PhD program. It is so exciting, although not surprising in the least, to watch the Muday lab continue to thrive five years later.

To my family and friends outside the lab (or in a lab in a different building), again

I cannot thank you all sufficiently in just one paragraph. To my brilliant BSGP classmates, we have grown a lot over the past five years, and it has always been wonderful and cathartic to get together and celebrate/commiserate, even if it happens less and less often now. It is great to see everyone being so busy and successful, and I wish you all the best as we each pursue our next challenge. Thank you to John G. and

Carol Jacob for welcoming me so warmly into your home and family when John and I started dating, and for all the excellent conversation, laughter and adventures that we have had since- not to mention the delicious food and wine! Columbus would not feel like home without you two (and Joey and Wrigley of course!), and now we are so fortunate to have a home in Waco too!

To my family, I don’t even know what to say. Thank you, Mom and Dad, for your endless love and support, and for giving me the confidence to move to Columbus by myself at 21, not knowing anyone, and then answering the phone when I called every night for the first three months. Thank you for your unwavering belief in the power of education, and for pushing us to relentlessly set and achieve our goals. Thank you for being such amazing role models and advocates; you are truly the best parents ever. I

love you both so much. To Theresa, it has been such a pleasure to watch you grow up and find your own work to be passionate about, and I can’t wait to see what you do next.

Thank you for sharing your grammatical expertise and statistical analyses, and for keeping me hip. I am so lucky to have you as a sister and a best friend, and I love you.

Thank you to Kebo for being our family’s best friend for 12 wonderful years, and the only mammal (furry or otherwise) who could always make me laugh, not matter what was wrong. We love and miss you. And finally, thank you a million times over and then some to John for being the best partner, teammate, roommate, confidant, chef, drinking buddy, sports commentator, IT guy and support system out there. I would not be here today without you, nor would I have made so many good memories along the way. You are an amazing scientist and I cannot wait to see what you do next. I love you; thank you for your endless love, help, and patience, it means the world. Can we get a dog now?

vii

Vita

2011-2015………………B.A. Biology (with Honors) and French, Wake Forest University

2015-present…….………………... Graduate Research Fellow, The Ohio State University

Publications

1. Woods PS, Doolittle LM, Rosas LE, Nana-Sinkam SP, Tili E, Davis IC. Increased expression of microRNA-155-5p by alveolar type II cells contributes to development of lethal ARDS in H1N1 influenza A virus-infected mice. Virology. 2020;545:40-52.

2. Doolittle LM, Davis IC. Influenza in Smokers: More than Just a Cause of Symptom

Exacerbations? American Journal of Respiratory Cell and Molecular Biology.

2018;59(6):670-1.

3. González JF, Alberts H, Lee J, Doolittle L, Gunn JS. Biofilm Formation Protects

Salmonella from the Antibiotic Ciprofloxacin In Vitro and In Vivo in the Mouse Model of chronic Carriage. Scientific Reports. 2018;8(1).

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4. Woods PS, Doolittle LM, Hickman-Davis JM, Davis IC. ATP catabolism by tissue nonspecific alkaline phosphatase contributes to development of ARDS in influenza- infected mice. American Journal of Physiology-Lung Cellular and Molecular Physiology.

2018;314(1):L83-L92.

5. Woods PS*, Doolittle LM*, Rosas LE, Joseph LM, Calomeni EP, Davis IC. Lethal

H1N1 influenza A virus infection alters the murine alveolar type II cell surfactant lipidome. American Journal of Physiology - Lung Cellular and Molecular Physiology.

2016;311(6):L1160.

Fields of Study

Major Field: Biomedical Sciences Graduate Program

Table of Contents

Abstract ...... i

Dedication ...... iii

Acknowledgments ...... iv

Vita ...... viii

Table of Contents ...... x

List of Tables ...... xiv

List of Figures ...... xv

Chapter 1. Literature Review ...... 1

1.1 Influenza ...... 1

Influenza Biology ...... 1

Influenza Pathogenesis ...... 5

IAV Infection and Host Metabolism ...... 9

Influenza Epidemiology ...... 12

Influenza Prophylaxis and Treatment ...... 14

Mouse Models of Influenza Infection ...... 16

1.2 Acute Respiratory Distress Syndrome ...... 18

Clinical Characteristics ...... 18

Epidemiology ...... 22

Prognosis and Treatment ...... 24

Animal Models of ARDS ...... 26

1.2.4.1 Virus-Induced ARDS ...... 27

1.2.4.2 Other Models of ARDS ...... 28

1.3 Alveolar Type II Cells ...... 29

Alveolar Type II Cells in the Normal Lung ...... 29

ATII Cell Bioenergetics ...... 33

ATII Cells in the Diseased Lung ...... 34

Role in Influenza Pathogenesis ...... 36

1.4 Mitochondria ...... 37

Mitochondrial Structure ...... 37

1.4.1.1 Mitochondrial Membrane Composition ...... 38

1.4.1.2 Phospholipid Biology and Synthesis ...... 39

1.4.1.3 Cardiolipin Biology and Synthesis ...... 42

1.4.1.4 Quality Control: Biogenesis and Mitophagy ...... 44

Mitochondrial Function ...... 47

1.4.2.1 Mitochondrial Energy Production ...... 47

1.4.2.2 Mitochondrial Signaling ...... 51

1.4.2.3 Mitochondrial Apoptosis ...... 53

Mitochondria in Viral Infection ...... 54

Mitochondria in Lung Disease ...... 58

1.5 Figures ...... 63

1.6 Tables ...... 73

1.7 References ...... 74

Chapter 2. Acutely Lethal Influenza Infection Alters ATII Cell Mitochondrial Structure and Function ...... 111

2.1 Abstract ...... 111

2.2 Introduction ...... 112

2.3 Materials and Methods ...... 116

2.4 Results ...... 126

2.5 Discussion ...... 132

2.6 Figures ...... 141

2.7 References ...... 148

Chapter 3. Acutely Lethal Influenza Infection Alters ATII Cell Bioenergetic Metabolism ...... 160

3.1 Abstract ...... 160

3.2 Introduction ...... 160

3.3 Materials and Methods ...... 162

3.4 Results ...... 168

3.5 Discussion ...... 171

3.6 Figures ...... 178

3.7 References ...... 182

Chapter 4. CDP-choline Rescue of Phospholipid Synthesis Improves Mitochondrial Function in Mice with IAV-induced ARDS ...... 190

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4.1 Abstract ...... 190

4.2 Introduction ...... 190

4.3 Materials and Methods ...... 193

4.4 Results ...... 198

4.5 Discussion ...... 200

4.6 Figures ...... 204

4.7 References ...... 208

Chapter 5. Future Directions ...... 216

5.1 Phospholipid Synthesis ...... 216

5.2 Fatty Acid Metabolism ...... 218

5.3 Mitochondrial Dysfunction ...... 219

5.4 CDP-choline ...... 220

5.5 Final Thoughts ...... 221

5.6 References ...... 222

References ...... 231

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List of Tables

Table 1.1: Berlin definition of ARDS...... 73

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List of Figures

Figure 1.1: Influenza A virus virion composition...... 63

Figure 1.2: Multiple RNA products are derived from a single IAV vRNA...... 64

Figure 1.3: Vectorial ion transport and alveolar fluid clearance...... 65

Figure 1.4: Synthesis, secretion and recycling of pulmonary surfactant...... 66

Figure 1.5: Mitochondrial structure and phospholipid composition ...... 67

Figure 1.6: Kennedy pathway de novo synthesis of phosphatidylcholine...... 68

Figure 1.7: CDP-DAG pathway de novo synthesis and remodeling of cardiolipin...... 69

Figure 1.8: An overview of cellular energy metabolism as it relates to mitochondria...... 70

Figure 1.9: Movement of protons and electrons through the electron transport chain complexes...... 71

Figure 1.10: Mitochondria play a role in multiple immune signaling pathways...... 72

Figure 2.1: ATII cell mitochondrial ultrastructure is altered by IAV infection...... 141

Figure 2.2: IAV infection alters mitochondrial ROS and mitochondrial mass in ATII cells...... 142

Figure 2.3: IAV infection alters cardiolipin synthesis and remodeling in ATII cells...... 143

Figure 2.4: IAV infection activates mitochondrial biogenesis in ATII cells...... 144

Figure 2.5: IAV infection reduces mitochondrial membrane potential in ATII cells but does not induce transition pore opening...... 145

Figure 2.6: IAV infection slows ATII cell oxygen consumption and alters mitochondrial energy production...... 146

Figure 2.7: IAV infection does not substantially alter electron transport chain gene expression in ATII cells...... 147

Figure 3.1: IAV infection alters ATII cell glucose metabolism but not glycolytic flux. .... 178

Figure 3.2: IAV infection alters TCA cycle flux in ATII cells ...... 179

Figure 3.3: IAV infection alters TCA cycle enzyme expression in ATII cells...... 180

Figure 3.4: IAV infection alters ATII cell ATP production rates...... 181

Figure 4.1: Daily CDP-choline treatment following IAV infection rescues ATII cell phosphatidylcholine (PC) synthesis...... 204

Figure 4.2: Daily CDP-choline treatment following IAV infection improves ATII cell mitochondrial physiology...... 205

Figure 4.3: Daily CDP-choline treatment improves ATII cell mitochondrial function. .... 206

Figure 4.4: CDP-choline treatment following IAV infection attenuates ARDS at 6 dpi. . 207

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Chapter 1. Literature Review

1.1 Influenza

Influenza Biology

Influenza viruses are segmented, single-stranded, negative sense RNA viruses in the Orthomyxoviridae family. They are further classified into four genera, influenza A,

B, C, and D viruses respectively, based on differences in nucleoprotein (NP) antigenicity.

Together, influenza A virus (IAV) and influenza B virus (IBV) are responsible for annual seasonal influenza epidemics and the majority of human influenza cases. Influenza C viruses can also infect humans, but they cause mild disease and cases are infrequent

(1). Recently identified influenza D viruses predominantly infect cattle and swine; human cases have yet to be reported (1). This overview of influenza biology will focus on IAV, as it causes significant morbidity and mortality during annual influenza epidemics, and possesses significant demonstrated pandemic potential (2, 3).

The IAV genome is composed of eight viral RNA (vRNA) segments (segment 1-

8) that encode proteins with roles in viral replication, assembly, budding, cell entry, and host evasion. The viral RNA-dependent RNA polymerase is composed of three subunits,

PA, PB1 and PB2. PA (segment 3) is responsible for IAV’s “cap snatching” endonuclease activity, as it cleaves capped host RNAs for replication of vRNA. PB2

(segment 1) binds and initiates vRNA transcription onto the hijacked cap structure, while

PB1 (segment 2) has polymerase activity and catalyzes vRNA elongation [reviewed in

(1, 4, 5)]. vRNA segments are encapsulated by nucleoprotein (NP; segment 5) and matrix (M1; segment 7) proteins which mediate nuclear trafficking of vRNA, as well as virion assembly (1, 4). Neuraminidase (NA; segment 6) and membrane protein (M2;

segment 7) coordinate viral budding and release from the cell (1, 4). Receptor binding and cell entry is directed by hemagglutinin (HA; segment 4) and M2. IAV also encodes two non-structural proteins (NS1 and NS2; segment 8). NS1 antagonizes the host cell- induced interferon (IFN) response to IAV, and NS2 coordinates nuclear export of vRNA and associated NP, referred to as viral ribonucleoprotein (vRNP) complexes (1, 4).

Splicing of vRNA, regulated by NS1, allows IAV to synthesize additional proteins from the eight genome segments, including PB2-S1 (segment 1), PB1-F2 (segment 2), and

PA-X (segment 3). Each of these accessory proteins plays a role in modulating and evading the host immune response to IAV infection (1). The composition of the influenza virion is illustrated in Figure 1.1.

The IAV life cycle [reviewed in (2, 4, 6)] begins with HA attachment to the virus’ cellular receptors, sialic acid residues on host cell glycoproteins with either an α2,3- or an α2,6- linkage, depending on virus strain and host species. HA binding triggers virion endocytosis via clathrin-mediated endocytosis or macropinocytosis, followed by trafficking to the endosome. Within the endosome, cellular proteases cleave HA into

HA1 and HA2 subunits. The low pH in the endosome facilitates fusion of the IAV envelope membrane with the endosomal membrane by inducing a conformational change in HA2 to expose a fusion peptide. Activation of M2 ion channel activity by the low endosomal pH results in acidification of the virion and the release of vRNPs, containing the viral genome, into the host cell cytoplasm.

Once released into the cytoplasm, IAV vRNPs (vRNA and NP complexes) must be trafficked to the nucleus for replication of the viral genome and transcription and

translation of viral proteins. vRNPs contain nuclear localization signals and use the importin-α-importin-β pathway to enter the nucleus. In addition to a segment of the IAV genome, each vRNP contains the viral polymerase (PA, PB1, PB2). Within the nucleus, replication of the viral genome by the IAV polymerase begins with transcription of complimentary viral RNA (cRNA) templates, followed by the transcription of vRNA copies from the cRNA. Temporally, transcription and translation of vRNAs within the cell occur before viral genome replication (6). A diagram showing the RNA products derived from vRNA segments is shown in Figure 1.2. Transcription of viral mRNAs from vRNA uses the viral polymerase but splicing and translation use cellular machinery. To mimic host mRNA, IAV polymerase uses cap snatching from host transcripts to generate primers for viral transcripts. Synthesis of a viral mRNA is terminated with polyadenylation triggered by a poly-U sequence at the 5’ end of each vRNA, resulting in mRNA that can be exported from the nucleus alongside host transcripts. As previously mentioned, during mRNA transcription IAV employs cellular splicing mechanisms to produce additional viral proteins, although this process is less efficient than for host mRNA (4).

IAV NS1 has been identified as a regulator of viral mRNA splicing as a function of its nuclear export role, which may regulate how many full length transcripts remain in the nucleus for long enough to facilitate splicing (4). Mature viral mRNAs are exported from the nucleus to the cytoplasm using host mRNA export mechanisms (7), and viral proteins are translated on ribosomes, either cytosolic (PB1, PB2, PA, NP, NS1, NS2 and

M1) or endoplasmic reticulum-associated (HA, NA and M2) (6).

Newly synthesized NP, PA, PB1 and PB2 are imported back into the nucleus, again utilizing the importin-α-importin-β pathway, to participate in viral replication and

gene transcription. NS1 is also imported into the nucleus where it targets multiple pathways to prevent IFN synthesis and attenuate the IFN response to IAV (3, 4). M1 and

NS2 are also imported into the nucleus to coordinate the nuclear export of vRNPs for virion assembly. M1 associates with vRNP complexes and prepares them for nuclear export, while NS2 recruits nuclear export machinery and interacts with M1 to direct vRNP export (4, 6). Once in the cytoplasm, vRNPs are translocated to the plasma membrane by Rab11 via interaction with PB2 (6). IAV buds from the apical membrane of polarized epithelial host cells at regions containing high levels of sphingolipids and cholesterol, termed lipid rafts. Following post-translational modification, HA, NA, and M2 traffic to viral assembly sites via apical sorting signals (4). HA and NA contain transmembrane domains that are preferentially incorporated into lipid rafts, while M2 binds cholesterol moieties on the cytoplasmic side of the cell membrane (4). Interactions between M1, NS2, HA and NA, as well as between M2 and vRNPs may play a role in the collection of virion components at plasma membrane lipid rafts for assembly. An infectious virion must contain all of the eight genome segments. It remains unclear how influenza viruses achieve this. One packaging model suggests random incorporation, resulting in only a small portion of virions containing the correct segments for infectivity

(4). A second model suggests selective incorporation, based on a unique packaging signal on each vRNA segment, resulting every virion produced being infectious. Growing experimental evidence supports this second model (4).

The final step in the IAV life cycle is budding and release from the host cell.

Budding requires induction of significant plasma membrane curvature, which is facilitated by HA, NA, M2, and results in membrane fusion at the base of the bud (4).

Membrane scission completes the formation of the viral envelope, which will remain attached to the host cell by HA binding to sialic acid residues. NA cleaves sialic acid to release the virus from the host cell and to prevent aggregation of nascent virions. Anti-

IAV therapeutics target NA activity to prevent viral infection and disease.

Influenza Pathogenesis

Influenza is transmitted primarily by inhalation of respiratory droplets and aerosols containing infectious IAV particles. The virus attaches to epithelial cells along the respiratory tract, and acute symptom onset occurs after a 1-2 day incubation period

(8). Symptoms are both systemic- fever, chills, headache, muscle pain, malaise and loss of appetite, and respiratory- cough, sore throat and runny nose. The typical duration of systemic symptoms is 3-8 days, but respiratory symptoms and malaise may last for up to two weeks (8). Infected individuals shed the most virus during the beginning of clinical illness (fever, cough, fatigue), but can also shed during the incubation phase (8).

Typically, IAV shedding is no longer detectable about a week after symptoms begin, coincident with the cessation of clinical illness (8).

IAV infection of respiratory epithelium activates the innate immune response, which attempts to slow or prevent viral replication. Activation of innate immunity is based on the detection of pathogen-associated molecular patterns (PAMPs) by pattern recognition receptors (PRRs) in infected cells. IAV produces several PAMPs, including double-stranded RNA (dsRNA), single-stranded RNA (ssRNA), and 5’-triphosphate

RNA. The PRR toll-like receptor (TLR) 3 recognizes dsRNA, while TLR7 and TLR8 recognize ssRNA. Retinoic acid-inducible gene I (RIG-I) is a PRR that recognizes 5’-

triphosphate RNA. Another group of PRRs, NOD-like receptors (NLRs) recognize other viral products (9). PRR activation initiates the secretion of inflammatory mediators such as leukotrienes, prostaglandins, cytokines, chemokines and type I IFNs. These molecules contribute to the clinical presentation of influenza, causing fever, malaise and inflammation, while also recruiting myeloid and lymphoid cells to infected tissue (9).

TLRs detect IAV-associated PAMPS primarily in acidified late endosomes upon viral entry. TLR3 is additionally expressed on the surface of epithelial cells of the lower respiratory tract (LRT), and, once activated, signals through TRIF (TIR-domain containing adaptor protein-inducing-IFN-b) to activate NF-kB and IRF (IFN regulatory factor) 3 to induce the production of Type I IFNs and pro-inflammatory cytokines (9). In comparison, TLR7 and TLR8 are expressed only in endosomes and signal through

MyD88 upon activation, activating NF-kB and multiple IRFs, resulting in expression of various pro-inflammatory cytokines and IFNs, depending on cell type (9). Overall, TLR activation is crucial for mounting an IFN response to IAV.

RIG-I is the major RLR (RIG-I-like receptor) involved in IAV PAMP sensing.

Constitutively expressed in epithelial cells, macrophages, and dendritic cells, RIG-I is located in the cytosol and binds ssRNA with 5’-triphosphates, preferentially binding short viral RNA transcripts. Upon activation, RIG-I binds ATP and complexes with the mitochondrial antiviral signaling protein (MAVS). MAVS activates NF-kB and IRF3 to induce type I IFNs and inflammatory cytokine production (9).

The inflammatory milieu in the respiratory tract induced by PRR signaling is characterized by high levels of pro-inflammatory cytokines such as TNF-a, IFN-g, IL-

1b, IL-6, and IL-17 (10). Chemokines, including IL-8, CXCL10 and CCL2, recruit inflammatory cells, predominantly neutrophils, but also macrophages, natural killer cells, and dendritic cells. The extent of neutrophil infiltration strongly correlates with influenza severity, possibly due to the release of factors from neutrophils that can damage lung tissue, such as neutrophil extracellular traps (NETs) (9). Dendritic cells are primarily involved in IAV antigen presentation to T cells during the development of an adaptive immune response to IAV infection, which takes at least 5 days to become fully active (9).

Although the adaptive response to IAV, via T cell maturation and antibody production, plays a major role in controlling IAV replication and preventing future infection with the same virus, it will not be discussed in detail here as the timeframe for this response is outside of acute IAV-induced illness, our area of interest.

Many of the complications associated with influenza are related to the immune response to the virus. Other aspects of severe disease are caused by direct viral infection and damage of the respiratory epithelium, compromising lung function. Infection of alveolar epithelial cells, which facilitate gas exchange, is a key driver of severe disease, as it brings viral and inflammatory damage to the functional units of the lung.

Alveolar infection can result in the loss of alveolar structure and epithelial barrier integrity, from IAV-induced cell death as well as from excess inflammation. The loss of alveolar structure reduces gas exchange capacity, which can lead to acute respiratory distress syndrome (ARDS) and respiratory failure (11).

Severe pneumonia can occur with influenza, either due to the viral infection or the development of a secondary bacterial infection. Bacterial pneumonia following

influenza infection is commonly due to Staphylococcus aureus or Streptococcus pneumoniae (11). Nosocomial infections leading to pneumonia following hospitalization for influenza may also occur (11). Viral and bacterial pneumonia associated with influenza are distinguishable based on their timelines. Viral pneumonia will occur coincident with clinical symptoms, while bacterial pneumonia often occurs one to three weeks after resolution of symptoms and may present as an influenza “relapse” (11).

Research suggests that damage to lung tissue caused by IAV infection may promote bacterial colonization of the respiratory tract by eliminating physical barriers to bacterial attachment (12). Additionally, mobilization of the anti-viral response may dampen the immune system’s ability to respond to bacterial pathogens for a period of time following

IAV infection, creating a window of opportunity for bacterial infection of the respiratory tract (11-13). Non-pulmonary complications of influenza include cardiac and neurological manifestations. Influenza can cause myocarditis and pericarditis and can exacerbate underlying pulmonary disease. The virus is also associated with multiple neurologic complications, including meningitis, aseptic encephalitis, and Guillain-Barré syndrome

(8).

Resolution of influenza is temporally associated with viral clearance and the cessation of acute inflammation. The development of an antibody response to the infecting IAV strain plays a major role in viral clearance. IL-10 (an anti-inflammatory cytokine produced by CD8+ T cells) and macrophage-mediated suppression of the neutrophil response contribute to the resolution of inflammation in the lung (13). In severe influenza cases, where diffuse alveolar damage is widespread, reduction of inflammation is followed by regeneration of the alveolar structure to restore gas

exchange capability. This process involves matrix deposition, re-establishment of the epithelial-endothelial barrier, and proliferation of progenitor cells to re-populate the alveolus (13). The role of different cell populations in recovery from alveolar insult will be covered at the end of this chapter.

IAV Infection and Host Metabolism

Patients with metabolic disorders such as obesity, Type I diabetes and Type II diabetes are at higher risk for developing severe influenza and complications from infection (14, 15). Recently, research into the host metabolic changes that may underly these risk factors has expanded greatly. It has been observed that serum from patients with H1N1 viral pneumonia has reduced concentrations of glucose, glutamine, and fatty acids compared to serum from H1N1-infected patients with mild disease (16). A recent retrospective study found that positron emission tomography (PET) scans of pediatric cancer patients with IAV infection had significantly increased glucose uptake in their lungs during active infection, compared to a follow-up PET scan 8 weeks later (17). In the same study, treatment of mice with BEZ235, a PI3K/mTOR inhibitor that inhibits glycolysis in vitro, before and during infection with 2009 pandemic H1N1 IAV significantly improved survival and lung function, while reducing viral titer (17). These findings support previous research dating back to the 1950s that IAV replication requires sufficient glucose supply [reviewed in (18)].

Infection of mice with a semi-lethal dose of IAV PR/8/34 (H1N1) significantly reduced pyruvate dehydrogenase (PDH) activity in lungs, heart, skeletal muscle, and livers (19). PDH links glycolysis to mitochondrial ATP production by converting pyruvate

to acetyl-coA for entry into the tricarboxylic acid (TCA cycle). This decrease in PDH activity was due to increased enzyme inactivation by PDH kinase 4 (PDK4) following IAV infection. Treatment of mice with diisopropylamine dichloroacetate (DADA) following IAV infection rescued PDH activity and increased ATP levels in lungs, heart, skeletal muscle, and liver. DADA treatment also significantly increased survival and reduced IAV replication in this model (14, 19). It is difficult to parse out the specific metabolic processes that are advantageous or disadvantageous for IAV pathogenesis in vivo, as modification of one pathway will often lead to the development of compensatory mechanisms to promote organismal survival. However, it is clear that IAV thrives under specific metabolic conditions, and that its ability to replicate is closely tied to glucose availability. Drugs that modify systemic metabolism have great potential as anti-influenza therapeutics as their risk of driving drug-resistant strain evolution is low (18).

Additionally, some therapeutic candidates are already approved for clinical use in other indications (19).

More information is available about the effect of IAV on target cells. IAV infection increases glycolysis and reduces ATP production in Madin-Darby canine kidney (MDCK) cells (20). Immortalized kidney epithelial cells, MDCK cells are highly permissive to IAV infection, but do not closely represent the pulmonary epithelial cells invaded by IAV during infection of a host. In normal human bronchial epithelial (NHBE) cells, IAV infection increased glycolysis, but simultaneously increased mitochondrial respiration as well (17). Interestingly, these metabolic changes were unique to IAV infection and could not be replicated with polyinosinic polycytidylic acid (PolyIC), a molecule that mimics viral infection to the immune system (17). This suggests that the observed metabolic

changes are driven at least in part by IAV, not solely by the innate immune response to the virus. IAV infection of NHBE cells was associated with a transient increase in c-Myc early in infection, and metabolic increases associated with IAV infection in this model were ablated by treatment with a c-Myc inhibitor (17). C-Myc is a transcription factor that is considered to be a master regulator of cellular metabolism and a driver of metabolic changes and glucose addiction in cancer (21).

Another master metabolic regulator, the highly conserved serine/threonine kinase mechanistic target of rapamycin (mTOR), has also been identified to modulate host cell metabolism late in IAV infection. mTOR complex (MTORC) 1 responds to nutrient levels to coordinate cell growth, proliferation and protein synthesis. In multiple cell lines, IAV activated mTORC1 to promote viral replication. During infection, viral proteins HA and

M2 promoted mTORC1 activation (22). IAV also activated mTORC2, which regulates cell survival and cytoskeletal dynamics, in vitro. The mechanism for this is unclear, but it may involve viral protein NS1, which is known to modulate apoptosis (22). It has been proposed, that together these master regulators of cell state promote a pro-IAV environment at the end of the viral life cycle in stressed cells (22).

IAV infection also targets other metabolic pathways. Infection of mice with acutely lethal H1N1 IAV reduced levels of major glycerophospholipids, including phosphatidylcholine (PC) and phosphatidylethanolamine (PE) in alveolar type II epithelial cells, while cholesterol, sphingomyelin and diacylglycerol levels increased (23).

Infection of ferrets with the pandemic 1918 H1N1 virus resulted in increased PC and PE levels in lung and tracheal tissue (24). Cui et al. also identified changes in phospholipid

catabolism, sphingolipid metabolism and β-oxidation of fatty acids in whole lung tissue following IAV infection of mice with a sub-lethal dose of PR/8 (25). In another study, infection of mice with pandemic 2009 H1N1 altered glycerophospholipid metabolism in whole lung tissue, and this change was computationally associated with IFN-γ production (26).

Influenza Epidemiology

Seasonal influenza epidemics, such as those that occur each winter in temperate climates, are associated with significant morbidity and mortality. In the United States, seasonal influenza is associated with 140-960,000 hospitalizations and 12-79,000 deaths, depending the severity of the epidemic (15). The World Health Organization estimates that 3-5 million people are hospitalized with influenza each year, leading to between 250,000 and 500,000 deaths (27). Influenza pandemics can exacerbate morbidity and mortality, as exemplified by the “Spanish Flu” pandemic of 1918-19. This highly lethal H1N1 IAV infected an estimated one third of the world’s population, about

500 million people, and caused between 50 and 100 million deaths (3). The most recent influenza pandemic, in 2009-2010, was caused by a novel H1N1 virus, resulting in an estimated 60 million cases, 275,000 hospitalizations and 12,500 deaths (28). This IAV strain continues to circulate and cause illness during seasonal influenza epidemics.

A number of factors put people at increased risk for morbidity and mortality from influenza infection. Underlying respiratory and cardiovascular disease, diabetes mellitus, immunosuppression, and pregnancy can all enhance disease. The very young and the elderly are also at higher risk for severe illness and death from seasonal influenza (12),

although paradoxically this trend is often reversed for pandemic strains, with young adults suffering more severe illness from these viruses (3). During the 2009 influenza pandemic, a single-nucleotide polymorphism in the IFITM3 (interferon-inducible transmembrane 3) gene was identified as a predisposing factor for increased disease severity (29).

IAVs are identified and categorized by their antigenic surface glycoproteins, HA and NA. There are 16 known HA subtypes and nine known NA subtypes. H1N1, H2N2 and H3N2 viruses have all circulated in the human population in the past (30); currently

H1N1 and H3N2 viruses are co-circulating (31). Variation occurs even within IAV subtypes, a phenomenon referred to as antigenic drift. IAV encodes an RNA-dependent

RNA polymerase, which is error prone and lacks proof-reading capability, resulting in high mutation rates in the IAV genome. Mutation to the antigenic regions of HA and NA can result in a selective advantage by allowing the virus to evade antigenic detection and existing immunity to other strains. Antigenic drift necessitates yearly vaccination for circulating influenza strains and explains why humans experience clinical illness from

IAV multiple times over their lifespan.

Another genetic phenomenon that contributes to the pathogenicity of influenza viruses is antigenic shift. The IAV genome is composed of eight independent segments, which can re-combine with gene segments from other influenza viruses when a host cell is co-infected with two distinct strains. Antigenic shift results in the creation of novel

IAVs, containing gene segments from two “parent” viruses. The majority of the human

population will not have immunity to these novel viruses, and their pandemic potential is high, especially if they are able to spread easily from person to person (8).

Wild aquatic birds such as ducks, geese and swans are the reservoir host of IAV.

The virus infects intestinal epithelial cells in these species, causing minimal clinical disease. Antigenic shift is common in this setting, and multiple instances of IAV host- switching from birds to humans or other mammals have been documented (2). Notable species that are susceptible to IAV infection are domestic birds such as turkeys and chickens, and domestic swine. IAVs in these species have significant potential to cross- over and infect humans, and populations are closely monitored for novel viruses.

Several avian IAVs have caused outbreaks in humans, predominantly H5N1, as well as other avian H7 and H9 subtype IAVs. These viruses can be highly lethal to humans, although so far human to human transmission has been limited (1, 2).

Influenza Prophylaxis and Treatment

Vaccination against circulating influenza viruses is the main strategy employed to prevent morbidity and mortality from influenza. The dual goal of vaccination is to prevent both individual infection and community spread of the virus, which is highly contagious.

Immunization against circulating influenza viruses must be done yearly, as antigenic shift and drift are constantly producing virus strains that are antigenically novel. The World

Health Organization issues annual recommendations on which strains should be included in the next year’s vaccine, based on expert analysis and predictions (32). Most influenza vaccines are inactivated and contain purified viral protein components administered by intramuscular injection to generate a humoral immune response (33).

Trivalent vaccines contain antigens from H1N1 and H3N2 IAV stains alongside the dominant circulating IBV lineage. Quadrivalent vaccines contain antigens from both circulating IBV lineages (8). Unfortunately, influenza vaccine uptake remains poor in the

United States, with an estimated overall uptake of 47% in the 2014-2015 flu season (8).

Once infected, treatment of influenza is centered around antiviral medications that can reduce viral replication and duration of symptoms. For maximal effectiveness, antiviral treatment must be initiated as soon as possible after onset of symptoms.

Oseltamivir, a NA inhibitor, is the preferred anti-influenza medication (31). NA inhibitors are active against both IAV and IBV strains. Other FDA-approved NA inhibitors include zanamivir (inhaled) and peramivir (intravenous). Cap-dependent endonucleases, such as baloxavir marboxil, are also active against both IAV and IBV. The adamantanes, amantadine and rimantadine, target IAV’s M2 ion channel, but these drugs are becoming obsolete as circulating IAV strains have developed widespread resistance (15, 31).

The approval of oseltamivir for influenza treatment was somewhat controversial, as the scientific community has perceived bias in the reporting of clinical trial outcomes

(34). A recent systematic review of full clinical study reports found that in uncomplicated influenza, oseltamivir reduced symptom duration by only 16.8 hours in adults (34). When used for prophylaxis, oseltamivir reduced infection by 55% overall, and by up to 80% within households. Studies of oseltamivir efficacy in severe influenza were not conducted for FDA approval of the drug (31), but post-marketing studies have found a mortality risk reduction of up to 35% in patients hospitalized with severe influenza (15).

Overall, early administration of oseltamivir is key for maximizing drug efficacy (15, 31),

and a therapeutic window remains for treatments that are more effective in attenuating influenza-induced disease.

Mouse Models of Influenza Infection

Murine models of influenza infection are commonly used to study the virus and its mechanisms. Advantages of working with mice include their low cost and minimal husbandry requirements compared to other animal models of influenza. Additionally, laboratory mice are a highly tractable genetic tool, with numerous strains available to assess the contribution of individual genes to influenza pathogenesis. Numerous parameters of influenza pathogenesis can be measured in live mice, including oxygen saturation, activity level, lung function, and weight loss. Additional parameters can be measured following euthanasia, including lung weight, lung pathology, and viral titers

(35). Survival rates following IAV infection can also be determined via mortality studies in mice, but it is important to note that death may be due to excessive weight loss following infection, not disease progression (36).

Although mice are not a natural host for IAVs, a selection of strains are able to successfully infect mice and induce respiratory disease. These include Influenza

A/Puerto Rico/8/34 (PR8), and Influenza A/WSN/33 (WSN), which were passaged for adaptation to mice. Interestingly, several highly pathogenic IAVs, including the 1918 pandemic H1N1, the 2009 pandemic H1N1, and highly pathogenic avian H5N1, can infect mice without prior adaptation (35). The success or failure of various IAV strains to colonize the murine respiratory tract may be linked to the sialic acid-linkage preference of HA. While humans express a-2,6 sialic acid linkages on upper respiratory tract (URT-

nose, nasal cavity, mouth, throat, larynx) epithelium and a-2,3 sialic acid linkages on lower respiratory tract (LRT- trachea, bronchi, bronchioles, alveoli) epithelium, mice express only a-2,3 linked sialic acids (35, 37). Strains that are able to infect cells with a-

2,3 linked sialic acids are often more pathogenic in humans due to their ability to colonize the LRT.

Although select IAV strains can induce lung damage in infected mice, often through LRT infection and viral pneumonia (35), influenza manifests differently in mice than in human patients. Mice do not exhibit clinical signs correlated to influenza in humans; they do not sneeze, cough or develop a fever (38). Instead IAV-induced clinical signs in mice include hypothermia, huddling, fur ruffling, and weight loss due to anorexia and dehydration. Additionally, while influenza is a highly contagious disease in humans, there is little concrete evidence that infected mice can transmit the virus to other mice.

Although evidence of transmission has been published, it is often circumstantial, and the consensus is that IAV transmission between mice is rare and inefficient (35).

A final limitation to mouse models of influenza is that common laboratory mouse strains lack functional MX1, the murine homologue of MXA, an interferon-induced gene that is a potent inhibitor of IAV in humans (39). The loss of this restriction factor facilitates IAV infection in mice, and wild mice and MX1-corrected laboratory strains are highly resistant to IAV (35). These limitations may reduce the direct application of results from mouse models to human influenza patients. However, murine infection models are able to recapitulate a number of the clinical signs of severe influenza and ARDS

experienced by critically ill patients (40), and research in murine models is critical for developing insights into caring for this population and testing therapeutic efficacy.

1.2 Acute Respiratory Distress Syndrome

Clinical Characteristics

Early reports of ARDS symptoms were linked to traumatic combat injuries that were initially stabilized, but progressed to respiratory failure within days (41). Referred to as “shock lung” and “wet lung” during World War I and World War II (41), this condition became more prominent during the Vietnam War as technological advances meant more soldiers survived initial battlefield trauma and progressed to develop “DaNang lung”.

Now known as ARDS, this syndrome was first defined as a clinical entity in the 1960s in civilian patient cohorts that were refractory to ventilator support. Like soldiers with

DaNang lung, these patients presented with hypoxemia, diffuse pulmonary infiltrates, alveolar collapse, and hyaline membranes, and had high mortality rates. The first advance in treating ARDS patients was made at Colorado General Hospital around this time, with the discovery of positive end-expiratory pressure (PEEP) (42). The first description of ARDS, and the improvement in ventilation success and patient survival with the use of PEEP, was published by The Lancet in 1967 (43).

Since then, the definition of ARDS has been refined multiple times. The two most recent definitions are the American-European Consensus Conference (AECC) definition of 1994, and the current Berlin definition, proposed in 2012. Many aspects of the AECC definition are still used in clinical practice and ARDS research (44). Of relevance here is the inclusion of acute lung injury (ALI) in the AECC definition. ALI is an overarching term

for acute respiratory failure that may be associated with less severe hypoxemia than

ARDS (45). Although the Berlin definition no longer recognizes ALI as a clinical entity, the categorization continues to be used in respiratory research to describe mild lung injury or respiratory failure where ARDS diagnostic criteria cannot be measured.

The Berlin definition of ARDS incorporates multiple clinical findings and parameters. For acute respiratory failure to be considered ARDS, it must occur within one week of a known clinical insult, or new or worsening respiratory symptoms. Bilateral opacities must be observed on chest radiograph or CT scan. Pulmonary edema in ARDS must be independent of cardiac failure or fluid overload. Finally, the Berlin definition subdivides ARDS into three categories based on severity of the defining clinical feature for this syndrome- oxygenation measured by PaO2/FiO2. This ratio measures hypoxemia by quantifying how much inspired oxygen (FiO2) is able to cross the alveolar barrier into the blood stream (PaO2). In a healthy lung, the PaO2/FiO2 is 600 mmHg or greater (40).

Mild ARDS occurs when a patient’s PaO2/FiO2 is between 300 and 200 mmHg. Moderate

ARDS is between 200 and 100 mmHg, and a PaO2/FiO2 below 100 mmHg is considered severe ARDS (44). Table 1 summarizes the components of the Berlin definition of

ARDS.

As a clinical syndrome that leads to acute respiratory failure, ARDS has diverse causes, all of which lead to the development of pulmonary edema, significant hypoxemia, and reduced lung compliance. Many ARDS cases are associated with inflammatory injury to the lungs, and the two most common causes are bacterial and viral pneumonia (46). Non-pulmonary sepsis is another common infectious cause. Non-

infectious clinical events such as severe trauma, aspiration of gastric contents, pancreatitis and adverse drug reactions can also cause ARDS (47). These conditions all induce the pathophysiologic mechanisms underlying ARDS development, including a dysregulated inflammatory response, increased epithelial and endothelial permeability in the lung, and impaired alveolar fluid clearance.

In line with the infectious causes of ARDS, a maladaptive innate immune response, driven by PRR activation on lung epithelium and alveolar macrophages, drives excessive pulmonary inflammation, which can initiate ARDS. Development of a pro-inflammatory environment in the lungs includes the production of chemokines and cytokines that recruit neutrophils to the lung. Neutrophil recruitment can neutralize pathogens, but the release of proteases, reactive oxgen species (ROS) and NETs can also damage alveolar tissue (46, 48). The extent of neutrophil infiltration in the lung is associated with increased severity and mortality in ARDS (48). Increased neutrophil presence in the lung is also associated with increased influenza severity (49).

Alongside increased inflammation, increased endothelial and epithelial permeability also contribute to the development of ARDS. Lung injury and pro- inflammatory signaling molecules disrupt vascular endothelial cadherin junctions that are required to maintain barrier integrity in pulmonary microvessels. The destruction of these junctions increases endothelial permeability, and results in fluid accumulation in the alveolar space (46, 50). Inflammatory processes also disrupt epithelial junctions, which are much less permeable than endothelial junctions under homeostatic conditions. In

ARDS, multiple pathologic processes can contribute to destruction of the epithelial

barrier, including immune cell migration and direct killing of alveolar epithelial cells by infectious agents. In combination with increased endothelial permeability, damage to the epithelial barrier leads to pulmonary edema driven by fluid accumulation in the alveolar space (46). It is important to note that in ARDS, this increased permeability is the main driver of pulmonary edema, as opposed to increased pulmonary vascular pressure during left heart failure, which causes cardiogenic pulmonary edema (46, 47).

Fluid is normally removed from the lung parenchyma by alveolar fluid clearance

(AFC), a process by which alveolar epithelial cells generate an osmotic gradient that favors the movement of water across the epithelium and into the interstitial space, as illustrated in Figure 1.3. In ARDS, the epithelial sodium channel (ENaC) and the sodium/potassium ATPase pump (Na/K ATPase) that drive fluid clearance are both inhibited, resulting in decreased AFC (47). Multiple elements of the ARDS disease state have been shown to contribute to reduced AFC, including hypoxia, hypercapnia, biomechanical stress (such as that induced by mechanical ventilation), and pro- inflammatory cytokines (47). AFC is the major mechanism for resolving pulmonary edema, and as such is critical for resolution of ARDS. ARDS patients with impaired AFC have decreased survival rates (46, 47). Other processes that help to restore alveolar structure also contribute to ARDS resolution and recovery of normal gas exchange.

These include proliferation and migration of alveolar epithelial cells and pulmonary fibroblasts to repair the epithelial and endothelial barriers and restore normal permeability (51).

A major recent advance in ARDS research is the use of large data sets from

ARDS clinical trials to identify sub-phenotypes of ARDS. One subtype, referred to as

“uninflamed” or “hypoinflammatory” is associated with low plasma levels of biomarkers for inflammation, coagulation, and endothelial activation, as well as a lower mortality rate, around 15% (52). This subtype is also associated with upregulation of mitogen- activated protein kinase (MAPK) signaling pathways (53). The second subtype, referred to as “reactive” or “hyperinflammatory” ARDS, is associated with high plasma levels of the same biomarkers, as well as high levels of neutrophil activation, and upregulation of oxidative phosphorylation and cholesterol metabolism pathways in blood leukocytes (52,

53). Patients with reactive ARDS have much higher mortality rates than uninflamed

ARDS patients, around 40% (52, 53).

Epidemiology

An estimated 200,000 patients are diagnosed with ARDS each year in the United

States, resulting in approximately 75,000 deaths annually (54). It is difficult to gauge the global prevalence of ARDS as diagnosis of this condition is closely linked to the availability of ICU beds and ventilatory support, which varies greatly between countries

(55). The LUNG-SAFE study, which gathered data on the international burden of ARDS, reported that 10% of ICU patients worldwide have ARDS, as categorized by the Berlin definition. Additionally, 23% of mechanically ventilated patients have ARDS. Overall,

40% of ARDS patients will die during hospitalization (55, 56). Post-hospitalization, mortality from ARDS remains high, with studies reporting 28/30 day mortality of 30% and

60 day mortality of 32% (57).

It is important to note that these statistics may underestimate the burden of

ARDS. This syndrome is often underdiagnosed. The LUNG-SAFE study found that clinicians missed 40% of ARDS cases, in spite of specific training on diagnosing ARDS

(56). Additionally, a small cohort study from Wuhan, China found that 41.8% of hospitalized coronavirus disease 2019 (COVID-19) patients had ARDS (58). It is probable that the COVID-19 pandemic will inflate ARDS incidence statistics for years to come. Large cohort studies have shown that 35-50% of ARDS cases are secondary to viral or bacterial pneumonia. Non-pulmonary sepsis is the cause in 30% of cases, while aspiration and trauma each contribute 10% of ARDS cases (55).

A number of non-acute factors can increase an individual’s ARDS risk. Age, non- white race, smoking, alcohol abuse, and ozone exposure have all be identified as risk factors (55, 56). The LUNG-SAFE study found that women are less likely to develop

ARDS than men, possibly due to protective effects of estrogen in the lung (56).

Interestingly, several studies have shown that obesity and diabetes may reduce ARDS risk (55, 56). Among ARDS patients, increased hospital mortality is associated with advanced age, active neoplasm, and chronic liver disease (55). Overall, ARDS presents significant risk to vulnerable populations and a substantial burden to intensive care units

(ICUs).

In addition to high levels of mortality, ARDS is also associated with substantial morbidity. At five years after ICU discharge, ARDS patients report a 25% reduction in physical function and a 17.5% reduction in overall health, compared to the general population. Diffusing capacity, a measurement of lung function, remains at or below the

lower limit of normality for ARDS survivors after one year. Six minute walk test distance, a measure of global physical function, also remains below the lower limit of normality

(59). Mental health issues are also common in ARDS survivors; 43.5% experience post- traumatic stress disorder (PTSD) at the time of hospital discharge, and 28% of survivors still have PTSD at five years (59). Incidence of PTSD is related to durations of mechanical ventilation, sedation and ICU stay. Other neuropsychological sequelae include depression (50% of survivors at one year, 58% at two years), long term cognitive impairment and executive dysfunction (55% of survivors at two years). Even after hospital discharge, ARDS survivors experience high levels of morbidity that constitute an additional medical, social and economic burden.

Prognosis and Treatment

To date, no therapeutics have been developed that ameliorate ARDS. Instead,

ARDS is treated with supportive care, centered around mechanical ventilation to maintain sufficient gas exchange while resting the respiratory muscles (54). A major advance in ARDS care was the recognition of ventilator induced lung injury (VILI), which is a result of additional stress and strain placed on lung tissue during mechanical ventilation (60). VILI is induced by volutrauma, large tidal volume during breathing, combined with atelectrauma, or repetitive opening/closing of distal airways and alveoli.

Together, these processes trigger the release of inflammatory mediators that contribute to organ dysfunction (61). To prevent VILI, modern ventilation strategies use a relatively high respiratory rate, a low tidal volume of around 6 mL/kg of predicted body weight, plateau pressure below 30 cmH2O, and high PEEP for patients with moderate to severe

ARDS (60). This ventilation strategy is considered to be protective in ARDS. Other components of mechanical ventilation may include neuromuscular blockade and prone positioning. Neuromuscular blockade is used to prevent spontaneous breathing in moderate to severe ARDS (60). Spontaneous breathing can contribute to VILI and increase mortality (60, 61). Prone positioning is used to promote homogeneous lung inflation for a more uniform distribution of mechanical force in moderate to severe ARDS

(54). Prone position should be maintained for at least 16 consecutive hours for maximum survival benefit (60). Neither neuromuscular blockade nor prone positioning have beneficial effects in mild ARDS. It is critical that all supportive care measures, including ventilation parameters be re-evaluated every 24 hours to assess safety and efficacy for the patient (60).

Additional supportive care techniques may provide benefit in ARDS, especially severe cases, but these interventions are supported by less evidence and are therefore still considered experimental. Extracorporeal CO2 removal (ECCO2R), uses an extracorporeal gas exchanger to remove CO2 from the blood, supporting ultraprotective ventilation by reducing CO2 removal in the lungs. A small clinical trial indicated that

ECCO2R may increase ventilator-free days in hospitalized patients with severe ARDS

(54, 62). A similar technique, extracorporeal membrane oxygenation (ECMO), has become popular since it was first used in ARDS patients during the IAV H1N1 pandemic of 2009. ECMO is recommended as a “last resort” strategy for severe ARDS patients because of mixed clinical data regarding its benefit, although it is associated with minimal adverse events (60). Extracorporeal gas exchange is a complex medical technique that is only available in highly specialized care settings.

Lung recruitment maneuvers during mechanical ventilation use increased airway pressure to open collapsed lung parenchyma, with the goal of increasing oxygenation.

This approach has generated mixed results in clinical research, with some studies indicating a survival benefit while others find increased mortality in the lung recruitment group (54). Each of these treatment strategies continues to be evaluated in clinical research. By their nature, mechanical ventilation and associated therapies require constant monitoring by highly trained medical staff, which is only possible in an ICU setting. None of these strategies are scalable to cope with the ARDS case burden that could occur during a severe influenza epidemic or a pandemic.

In part to address this deficiency, a number of pharmacologic strategies have been tested for ARDS, with minimal success. b2 agonists, which can increase AFC, successfully reduced pulmonary edema in a Phase 2 trial, but showed increased mortality in subsequent trials. Keratinocyte growth factor (KGF), an important compound for alveolar epithelial repair, also increased mortality, as well as duration of mechanical ventilation, in clinical trials. Treatment with anti-inflammatory statins showed no difference from the placebo group in multiple studies (54). Other drugs that have been tested with mixed or negative results include corticosteroids, diuretics, macrolide antibiotics, and synthetic pulmonary surfactants (63-65).

Animal Models of ARDS

ARDS is a complex medical condition, which can be partially, but not fully, recapitulated in animal models. While ideal for conducting the hypothesis-driven research necessary to better understand this devastating syndrome, laboratory animal

species lack the range of pre-existing conditions, exposures and risk factors that underly

ARDS in many human patients (66). Additionally, lung structure, function and inflammatory processes will vary between humans and animal species, so results cannot always be extrapolated to higher species or patient care (67). Large animal models of experimentally-induced ARDS, such as sheep, dogs, cats and pigs are considered to be more comparable to human ARDS, but these models are costly and technically challenging (68, 69). Mice, in comparison, are more cost-effective and experimentally accessible, in terms of reagents, techniques and genetic modification. Mice are the predominant animal model used in ARDS research (67). ARDS can be experimentally induced in mice a number of ways.

1.2.4.1 Virus-Induced ARDS

Mouse-adapted IAVs are a well-established “one hit” model for inducing ARDS that has been used by multiple research groups [reviewed in (67)]. IAV is also the leading viral cause of ARDS in humans (11). Intranasal inoculation of mice with a high dose of mouse-adapted influenza A/WSN/33 induces clinical signs consistent with ARDS in humans in a matter of days, with minimal biological variability (40, 70, 71). This includes the acute onset of hypoxemia, pulmonary edema, reduced lung compliance, and PaO2/FiO2 < 300 mmHg (40). Coronaviruses have also been used to induce ARDS in mice. Mice with a chimeric dipeptidyl peptidase 4 (DPP4) receptor can be inoculated with Middle Eastern respiratory syndrome coronavirus (MERS-CoV) and sustain viral replication. In this model, mice exhibited reduced pulmonary function and alveolar hemorrhage, which often occur in ARDS (72). In aged mice, recombinant infectious

clones of human severe acute respiratory syndrome coronavirus (SARS-CoV-1) induce respiratory distress and inflammatory profiles consistent with ARDS (73).

1.2.4.2 Other Models of ARDS

In reflection of the diverse clinical insults that lead to ARDS in humans, many experimental stimuli are used to induce ARDS in mice. Bacterial infection models are often used, as sepsis is a major cause of ARDS (74). Instillation of live bacteria or endotoxin into the lungs of mice elicits an inflammatory response and tissue damage consistent with ARDS (69). Models of extrapulmonary sepsis include intravenous delivery of bacteria and experimental peritonitis. Peritonitis is a common cause of sepsis in humans and is induced in mice by cecal ligation and puncture, which causes bacterial leakage from the gastrointestinal tract into the peritoneal space. Introduction of bacteria into the peritoneum on a carrier such as a sponge will also induce peritonitis.

Extrapulmonary sepsis models usually result in mild ARDS, with high biological variability (75).

VILI-induced ARDS is a commonly used model of sterile ARDS involving ventilation of mice with high tidal volumes and low PEEP. VILI induces pulmonary tissue damage and apoptosis of alveolar epithelial and endothelial cells but does not consistently trigger an inflammatory response. Although this model is considered to be highly clinically relevant, it is also technically challenging, requiring special equipment and expertise to establish consistent VILI in mice (69, 75). Direct instillation of acid, such as hydrochloric acid, into the lungs is used as a model for ARDS induced by gastric acid aspiration. This model reliably triggers clinical signs of ARDS in mice and can be titrated

to induced ARDS without immediate mortality, allowing for examination of mechanisms underlying the resolution of ARDS, an area that has been consistently difficult to study in animal models (76).

1.3 Alveolar Type II Cells

Alveolar Type II Cells in the Normal Lung

Alveoli, the functional gas exchange units within the lung, account for 99% of its internal surface area (77). Each alveolus is lined with type I (ATI) and type II (ATII) alveolar epithelial cells. The exchange of oxygen for carbon dioxide is facilitated by ATI cells, which are large but extremely thin, and as a result extremely fragile (78). ATI cells cover about 95% of the alveolar surface, and the remaining 5% is covered by ATII cells

(79). ATII cells are significantly smaller than ATI cells and cuboidal, with a diameter of approximately 10 µm (77). Despite this major size discrepancy, ATII cells are more abundant than ATI cells, making up 15% of all lung cells as compared to 8% for ATI cells

(80). Together, ATI and ATII cells maintain the alveolar epithelial barrier. The alveolar epithelium is covered with alveolar lining fluid and pulmonary surfactant to maintain humidity and surface tension. Other cellular components of the alveolus include alveolar macrophages and fibroblasts.

While the inherent delicacy of ATI cells makes them difficult to study, ATII cells have emerged as multi-functional cells that are critical for maintaining alveolar homeostasis. In the alveoli, ATII cells mediate fluid clearance, pulmonary surfactant production, innate immune responses, and epithelial barrier repair. Alveolar fluid clearance (AFC) is an important physiological process for maintaining fluid balance in

the lung and has been studied extensively. ATII cells create an osmotic gradient through transepithelial ion transport, and water moves down the osmotic gradient, from the alveolar space into the interstitium. ATII cells express epithelial sodium channels (ENaC) on their apical surface and Na/K ATPase on their basolateral surface. Sodium enters the cells through ENaC and is pumped out into the microcirculation by the Na/K ATPase (81,

82). Other ions and channels are also involved generating the osmotic gradient for AFC, including non-selective cation channels, cyclic nucleotide-gated channels, and the cystic fibrosis transmembrane conductance regulator, which transports chloride (83). ATI cells also contribute to AFC, although their role is less well-defined and they may not be able to compensate for a loss of ATII function (84). Defects in AFC have been identified in the pathogenesis of lung infections as well as ARDS (47, 85). Figure 1.3 illustrates the major components of AFC as well as the structure of the alveolus.

Another major role of ATII cells is the synthesis, secretion and recycling of pulmonary surfactant. Pulmonary surfactant lines the alveolar space and reduces surface tension to prevent the alveolus from collapsing, allowing it to remain inflated during breathing and facilitate gas exchange. The essential role of pulmonary surfactant is illustrated by respiratory distress syndrome in preterm infants who cannot produce their own surfactant and thus experience respiratory failure at birth [reviewed in (86)].

Pulmonary surfactant is composed of 90% lipids and 10% proteins. The lipid fraction is predominantly composed of phospholipids (80-85%). The majority of phospholipids in pulmonary surfactant are phosphatidylcholine (PC), of which one species, dipalmitoylphosphatidylcholine (DPPC), is mainly responsible for surface tension- reducing properties of pulmonary surfactant. This is because DPPC is highly

compressible. Other phospholipids in surfactant have important interactions with surfactant proteins. There are four surfactant proteins (SPs), SP-A, SP-B, SP-C and SP-

D. SP-A and SP-D are collectin family proteins, involved in lung innate immune responses (87). SP-B and SP-C are critical for surfactant function. They coordinate lipid packing and spreading to stabilize surfactant lipids during breathing, as well as to enhance their surface tension-lowering properties (88). SP-C is only expressed by ATII cells and is often used to identify these cells. Both SP-B and SP-C are secreted with surfactant lipids from lamellar bodies, specialized organelles in ATII cells where surfactant components are assembled (89). SP-A and SP-D have been found in other epithelial tissues, emphasizing their primary role in innate immune defense (86).

Surfactant phospholipids are synthesized in the ER. The major phospholipid synthesis pathways will be discussed later in this chapter. It is estimated that 45% of

DPPC is produced by the Kennedy pathway. The remainder is generated by remodeling of other PC species by phospholipase A2 (PLA2), which hydrolyzes unsaturated fatty acids and re-acylates PC with saturated fatty acids to generate DPPC (90). This remodeling reaction is catalyzed by acyl CoA:lyso-phosphatidylcholine acyltransferase

(LPCAT1), which is highly expressed in ATII cells (90). Phospholipid synthesis requires a sufficient supply of fatty acids, which are generated either by uptake of circulating fatty acids, or by de novo synthesis from lactate in ATII cells (90). Once secreted from lamellar bodies, pulmonary surfactant takes on multiple structural conformations in the airspace. Secreted surfactant components combine with SP-A to form tubular myelin, which is the surfactant form responsible for reducing alveolar surface tension. Tubular myelin can be isolated from alveolar fluid in large aggregates, along with other lamellar

body-like surfactant structures. Large aggregate components can be remodeled into small aggregates, vesicles which contain surfactant components but do not reduce surface tension (90). Figure 1.4 illustrates the full life cycle of pulmonary surfactant, from synthesis to secretion to recycling.

ATII cells participate in the innate immune response to bacterial and viral pathogens by secreting SP-A and SP-D, producing cytokines and chemokines, and presenting antigens. SP-A and SP-D have been shown to bind a number of respiratory pathogens, including Haemophilus influenzae, Mycoplasma pneumoniae,

Mycobacterium tuberculosis, respiratory syncytial virus (RSV), herpes simplex virus

(HSV) and IAV (87). Both SP-A and SP-D can aggregate pathogens in the alveoli, as well as regulate inflammatory signaling by immune cells. Additionally, SP-A can increase phagocyte uptake and killing of bound pathogens, while SP-D has direct anti-microbial activity (87). ATII cells can also produce and secrete an array of pro-inflammatory signals including interleukins (IL-6, IL8), IFNs (Type I and Type III), and chemo- attractants (MCP-1, MIP-1a, GM-CSF) that recruit macrophages and monocytes to the alveoli (91, 92). The immunomodulatory secreteome of ATII cells may vary based on the pathogen detected. Finally, ATII in both mice and humans express class II major histocompatibility complex (MHC), and have been shown to present microbial antigens to CD4+ T cells during M. tuberculosis infection (93).

The proliferative capacity of ATII cells has long been recognized. Following alveolar insult, ATII cells proliferate to repopulate and repair the epithelial barrier, eventually transdifferentiating into ATI cells (94). ATII proliferation and barrier repair is

critical to the resolution of alveolar damage, such as that sustained during ARDS (51).

The basic paradigm for this process is that ATII cells have stem cell characteristics and are able to proliferate and differentiate to ATI cells in response to a number of signals

(95, 96). These include Wnt signaling, bone morphogenic protein (BMP) signaling, hepatocyte growth factor (HGF), fibroblast growth factor (FGF) 7 and 10, and epidermal growth factor receptor (EGFR) (97-102). ATII cell dysfunction is linked to the development of pulmonary fibrosis, and fibroblast signaling plays a role in ATIl-mediated alveolar repair (98).

ATII Cell Bioenergetics

In homeostasis, ATII cells generate energy from glucose-derived substrates via oxidative phosphorylation. Due to their high demand for energy, ATII cells have triple the mitochondrial volume of other lung cells (103). Additionally, because of their unique direct exposure to oxygen, all lung cells express a distinct isoform of ETC complex IV,

COX subunit IV-2, which promotes aerobic oxidative phosphorylation (104). ATII cells can also use other substrates for energy production under conditions of cell stress.

During hypoxia, primary ATII cells conserve ATP by reducing the rate of oxidative phosphorylation and energetically expensive processes (105). Hypoxia also reduces glycolytic rate in primary ATII cells, although multiple genes associated with glycolysis are upregulated (105). ATII cells can also use lactate as an energy substrate to maximize mitochondrial ATP generation (106). Glucose, however, is required for ATII cell proliferation, as is common in many other cell types (106). ATII cells rely on fatty acids to generate phospholipids for pulmonary surfactant. Recent work has also shown

that ATII cells use fatty acid oxidation for energy production, and that disruption of this pathway plays a role in the development of acute lung injury (107). We will review the increasingly recognized role of ATII cell mitochondrial dysfunction in the development of multiple pulmonary disease states later in this chapter. This may include contributions due to bioenergetic failure, defective mitochondrial quality control, or mitochondrial participation in immune and apoptotic signaling pathways that promote a pro- inflammatory environment.

ATII Cells in the Diseased Lung

As ATII cells are at the center of alveolar homeostasis, they are also at the center of alveolar disease. ATII cell dysfunction has been noted to drive or contribute to a number of lung diseases, including ARDS, pulmonary fibrosis, COPD and pulmonary adenocarcinoma (108). Although a complete discussion of ATII cell contributions to the pathogenesis of these disease states is beyond the scope of this review, we will discuss the major mechanisms by which ATII cells may drive disease.

As previously discussed, ARDS is a disease of complex etiology, with many contributing causes and risk factors. Regardless of origin, non-cardiogenic pulmonary edema is a major component of ARDS. ATII cells drive AFC, which is the major mechanism for clearing edema fluid from the lungs (47). However, AFC by ATII cells is frequently impaired during ARDS (40, 109). A number of stimuli can reduce AFC by reducing ENaC and Na/K ATPase function (47). An aberrantly potent inflammatory response is another major component of ARDS (110), and ATII cells are capable of

producing a number of pro-inflammatory cytokines and chemokines in response to pulmonary pathogens.

Pulmonary fibrosis is characterized by the accumulation of fibroblasts and extracellular matrix (ECM) components secondary to ATII cell injury. Pulmonary fibrosis progressively reduces lung function and can be fatal in a matter of years. A number of mechanisms of cellular injury may contribute, including genetic mutations, cigarette smoking, viral infection and asbestos exposure. Alveolar injury is followed by the initiation of pro-inflammatory and pro-fibrotic secretory cascades that lead to abnormal

ECM deposition and the development of fibrotic lesions [reviewed in (111)]. COPD is another inflammatory lung disease which often occurs secondary to cigarette smoking and is characterized by irreversible airway obstruction driven by chronic bronchitis and emphysema. Inflammation in COPD is driven by multiple cell types, including ATII cells which secrete cytokines and chemokines and produce ROS [reviewed in (112)].

Lung cancers are the leading cause of cancer deaths worldwide. Lung adenocarcinomas express ATII cell markers, and the popular “ATII” cell line A549 was derived from a lung cancer tumor resection (113, 114). Mutations have been identified in

ATII cells that lead to tumor development, including the activation of oncogenic K-Ras and the suppression of p53 (115, 116). As with COPD and pulmonary fibrosis, cigarette smoking is a major risk factor for the development of lung damage and may contribute to

ATII cell injury and a pro-inflammatory environment that promotes oncogenic transformation.

Role in Influenza Pathogenesis

ATII cells play a central role in influenza pathogenesis as the primary site of IAV replication within the distal lung. In both humans and laboratory mice, ATII cells express a2,3-linked sialic acids on cell surface receptors, which are bound by IAV for cell entry.

ATI cells do not express this sialic acid linkage (37). In humans, IAV will also bind and enter lung epithelial cells expressing a2,6-linked sialic acids, which are not expressed in mice (37, 117). This may be why certain IAV strains are not infectious in mouse models.

The 1918 pandemic IAV (H1N1) and the highly pathogenic avian (H5N1) IAVs can establish productive infections in mice, suggesting a connection between a2,3- tropism and disease severity, possibly mediated by ATII cell involvement.

As already described, once IAV enters an ATII cell it begins its replication cycle, and in the process activates anti-viral innate immune responses. IAV infection induces numerous changes in the ATII cell transcriptome, of both infected and uninfected

(“bystander”) ATII cells. All ATII cells, regardless of infection status, upregulate IFN signaling and other anti-viral pathways, while only directly infected ATII cells activate pro-apoptotic pathways (118, 119). IAV infection reduces AFC in ATII cells by inhibiting

ENaC expression and activity [reviewed in (120)]. IAV infection also alters the ATII cell metabolome, including differential synthesis of lipids involved in pulmonary surfactant

(23, 25). Expression of pulmonary surfactant proteins is also reduced during infection

(121, 122). All of these changes support a role for ATII cells in the development of ARDS during IAV infection. ATII cells participate in the activation of a pro-inflammatory lung environment, undergo apoptosis which contributes to epithelial barrier destruction, show

altered surfactant production which would reduce lung compliance, and have reduced

AFC, which promotes the development of pulmonary edema.

IAV is not the only respiratory virus that targets ATII cells and causes serious disease in humans. SARS-CoV-1 also infects ATII cells, using angiotensin converting enzyme-2 (ACE2) as its receptor (123). Unlike influenza, which typically begins with upper respiratory tract infection that progresses to lower respiratory tract and ATII cells, studies of SARS-CoV-1 in macaques suggest that ATII cells are the initial site of infection for this virus (124). The recently identified SARS-CoV-2 also uses ACE2 for cell entry, in conjunction with the serine protease TMPSSR2 (125). Interestingly, TMPSSR2 is also used by highly pathogenic IAV strains, including 2009 pandemic H1N1 and avian

H7N9 for HA cleavage in ATII cells (126). A third zoonotic coronavirus, MERS-CoV, also infects pulmonary epithelium, using a2,3- and a2,6- sialic acid linkages for binding and dipeptidyl peptidase 4 (DPP4) for cell entry (127). Although MERS-CoV infection of ATII cells specifically has not been reported, it seems highly likely as ATII cells express a2,3- linked sialic acids and the virus is associated with severe respiratory disease.

1.4 Mitochondria

Mitochondrial Structure

Mitochondria are composed of two phospholipid bilayers, which divide this organelle into two distinct compartments with unique characteristics. Figure 1.5 illustrates the structure of a mitochondrion. The outer mitochondrial membrane (OMM) delineates the mitochondrion from the rest of the cell and contains proteins that coordinate trafficking of molecules into or out of the mitochondrion, as well as signaling

proteins that communicate between the mitochondrion and other organelles. Between the OMM and the inner mitochondrial membrane (IMM) is the intermembrane space

(IMS). The IMS is an aqueous environment through which proteins and metabolites must travel to reach the IMM. Generation of ATP by the electron transport chain (ETC) involves the transfer of protons into the IMS to generate proton-motive force for ATP synthase. The IMS is also a site of substantial ROS generation, and contains proteins to buffer the redox state of this compartment (128). The IMM is folded into invaginations called cristae, which increase the IMM surface area and contain the enzyme complexes that make up the ETC. The IMM also contains transport proteins to import the components necessary for ATP synthesis by oxidative phosphorylation. These components are stored in the mitochondrial matrix, the inner mitochondrial compartment that is enclosed by the IMM. The mitochondrial matrix is an important hub for many mitochondrial functions. The tricarboxylic acid (TCA) cycle occurs in the matrix, bridging cytosolic metabolism and mitochondrial ATP production by providing reducing agents for the ETC. The mitochondrial matrix also houses mitochondrial DNA (mtDNA), which must be protected to maintain mitochondrial integrity. Finally, the matrix plays a role in multiple cellular signaling pathways including calcium signaling, metabolic signaling and apoptotic signaling.

1.4.1.1 Mitochondrial Membrane Composition

To facilitate their various functions, the OMM and IMM bilayers have unique phospholipid compositions. The most abundant phospholipid in the OMM is phosphatidylcholine (PC), which comprises 54% of the total membrane lipid content.

Phosphatidylethanolamine (PE) accounts for 29%, while phosphatidylinositol (PI)

accounts for 14% (129). Numerous other lipid and phospholipid species are present, accounting for the remaining 3%. These include phosphatidylserine (PS), phosphatidylglycerol (PG), cardiolipin (CL), phosphatidic acid (PA), lysophospholipids, sterols and sphingomyelin (130). In the IMM, PC and PE together make up about 74% of the total membrane lipid content, with slightly less PC (40%) and slightly more PE (34%) than the OMM. The IMM also contains significantly more CL, about 18% of the total membrane lipid content (129). The conical shape of CL is critical for the folding of the

IMM into cristae. The proportional phospholipid composition of the OMM and IMM bilayers is depicted in Figure 1.5.

1.4.1.2 Phospholipid Biology and Synthesis

The majority of mammalian phospholipids, with the exception of CL, are synthesized in the endoplasmic reticulum (ER) and transported throughout the cell for membrane incorporation. PC is a fluid, tubular phospholipid that self-organizes into a planar bilayer and is critical for the structure of cellular membranes. In mammals, PC is predominantly produced via the Kennedy pathway. This pathway is initiated by the conversion of dietary choline into phosphocholine by choline kinase. Phosphocholine is converted into CDP-choline using a cytidine triphosphate (CTP) molecule via

CTP:phosphocholine cytidylyltransferase. This is the rate-limiting step of the Kennedy pathway. Finally, PC is synthesized by combining CDP-choline with a diacylglycerol

(DAG) molecule via CDP-choline:1,2-DAG cholinephosphotransferase (131). Figure 1.6 illustrates the Kennedy pathway of PC synthesis. PC an also be synthesized from PE by sequential methylation. This reaction is carried out by phosphatidylethanolamine methyltransferase (PEMT), and occurs predominantly in the liver in mammals (130).

PE, the second most abundant phospholipid in mitochondrial membranes, has a hydrophilic ethanolamine head group and a conical shape. It is not bilayer forming, but instead will self-organize into hexagonal phases. Inclusion of PE in phospholipid bilayers provides membrane tension and facilitates membrane curvature (130). Like PC, PE is primarily synthesized by the Kennedy pathway. Dietary ethanolamine is converted to phosphoethanolamine by ethanolamine kinase. Phosphoethanolamine is converted to

CDP-ethanolamine by CTP:phosphoethanolamine cytidylyltransferase. CDP- ethanolamine:1,2-DAG ethanolamine phosphotransferase, which is the rate-limiting enzyme for PE synthesis, converts CDP-ethanolamine and DAG to PE (132). The

Kennedy pathway of PE synthesis occurs in the ER, but mitochondria can also synthesize PE locally from PS. This occurs on the IMM via phosphatidylserine decarboxylase (PSD). Synthesis of PE via PSD is essential for maintaining mitochondrial morphology, and genetic loss of PSD is embryonic lethal in mice (133). Kennedy pathway synthesis of PE cannot compensate for the loss of mitochondrial PE synthesis via PSD.

PS is present at low levels in mitochondrial membranes but is an essential substrate for mitochondrial PE synthesis. PS is synthesized in the mitochondria- associated ER membrane (MAM) from both PC and PE. PS synthase enzymes replace the polar head groups of PC and PE with L-serine to produce PS. PS synthase-1 removes choline from PC while PS synthase-2 removes ethanolamine from PE (130,

132).

PI is the third most abundant phospholipid in mitochondrial membranes and is enriched in the OMM. The role of PI in mitochondrial function is not clear, but in general

PI is the precursor of phosphoinositides, important cellular signaling molecules (130). PI is synthesized in the ER from CDP-DAG and inositol by the enzyme PI synthase (134).

Similar to PS, PG is also present at low levels in mitochondrial membranes, but it is a critical substrate for CL synthesis. Synthesis of PG begins with the creation of PG phosphate (PGP) from CDP-DAG and glycerol-3-phosphate (G3P), catalyzed by PGP synthase. Next, PGP is dephosphorylated to PG (135). In mitochondria, specifically the

IMM, this reaction is carried out by protein tyrosine phosphatase mitochondrion 1

(PTPMT1) (136). This reaction is also considered the initial step in CL synthesis from mitochondrial PG.

PA, the simplest phospholipid in terms of structure, is only present in mitochondrial membranes small amounts, but serves as an important precursor in the synthesis of other lipids (130). PA can be synthesized from multiple precursors, but the major pathway begins with the acylation of G3P by G3P acyl transferase (GPAT) to produce lysophosphatidic acid (LPA). Subsequent acylation of LPA produces PA. LPA can also be produced from dihydroxyacetone phosphate (DHAP), for conversion to PA

(137). PA can also be generated from the phosphorylation of DAG and the hydrolysis of phospholipids (130). The multitude of sources for PA reflect the position of this molecule as central to the synthesis of biologically important lipids.

1.4.1.3 Cardiolipin Biology and Synthesis

CL (cardiolipin) is a phospholipid that is synthesized exclusively in mitochondria and closely associated with ATP production by this organelle. The relationship between

CL and ATP-producing membranes is evolutionarily conserved. In both bacteria and eukaryotic cells, CL is uniquely associated with ATP-producing membranes (138). CL is enriched in the IMM, where oxidative phosphorylation occurs. Like PE, CL has a conical shape that forces membrane curvature when it is incorporated into phospholipid bilayers.

In the IMM, CL interacts with each enzyme complex in the ETC, and is required for the enzymatic activity and structural integrity of these complexes. Additionally, CL is essential for respiratory supercomplex formation- the physical association of enzyme complexes to increase ETC efficiency and reduce ROS production (139). CL is also involved in ATP synthase oligomerization, which is critical for stabilizing and maintaining cristae structure (140). An additional function of CL is acting as a proton trap in the IMM: minimizing pH changes in the intermembrane space from ETC activity and supplying protons to ATP synthase (139).

CL is also involved in non-energetic processes in mitochondria. During mitochondrial fission, dynamin-related protein 1 (Drp1) forms a ring-like structure around mitochondria, and interacts with CL to promote membrane constriction and mitochondrial division (139, 141). CL also interacts with mitochondrial fusion protein optic atrophy-1

(OPA1), which facilitates IMM fusion (142). In addition to fusion and fission, CL plays a role in intrinsic apoptosis, which is coordinated by mitochondria. Redistribution of CL from the IMM to OMM promotes cytochrome c release, a pro-apoptotic signal. During extrinsic apoptosis, OMM CL serves as an activating platform for caspase 8, which in

turn triggers a pro-apoptotic signaling cascade that results in OMM permeabilization

(138, 139).

CL is synthesized de novo via the cytidine-5’-diphosphate-1,2-diacyl-sn-glycerol

(CDP-DAG) pathway, which is also responsible for PA synthesis. This pathway begins with the transfer of an acyl group from acyl-CoA to G3P by GPAT, forming lysophosphatidic acid (LPA). LPA is again acylated, this time by 1-acyl-sn-glycerol-3- phosphate acyltransferase (G3PAT), to form PA. The subsequent condensation of PA and CTP by CDP-DAG synthase (CDS) forms CDP-DAG; this is considered the rate- limiting step of the CDP-DAG pathway. Next, the phosphatidyl group of CDP-DAG is transferred to G3P to generate phosphatidyl glycerol phosphate (PGP) via PGP synthase. PGP is then hydrolyzed by PTPMT1 to form PG. Finally, PG receives another phosphatidyl group from another CDP-DAG molecule to form nascent CL in a reaction catalyzed by CL synthase (CLS) (143, 144).

Initial synthesis of cardiolipin utilizes a variety of acyl chains, which are then remodeled to generate highly specific molecular species of CL, which include acyl chains that are tissue- and organism-specific. Mature cardiolipin includes symmetric linoleic acid (18:2) chains. The remodeling process begins with phospholipase A2

(PLA2), which cleaves an acyl chain from CL to generate monolysocardiolipin (MLCL).

Three distinct enzymes are then able to re-acylate CL. Acyl-CoA:lysocardiolipin acyltransferase 1 (ALCAT1) and MLCL acyltransferase 1 (MLCLAT1) both use acyl-CoA as the acyl chain donor. Tafazzin (TAZ) meanwhile, is a transacylase that removes an

acyl chain from other phospholipids, such as mitochondrial PC and PE, and transfers it to CL (130, 145). Figure 1.7 illustrates de novo CL synthesis via the CDP-DAG pathway.

The physiological importance of CL is highlighted by the identification of defects in CL synthesis, remodeling, and integrity in multiple disease states including myocardial ischemia-reperfusion, heart failure, diabetes, and Parkinson’s disease [reviewed in (139,

145)]. The most prominent example of this is Barth syndrome, a rare, X-linked genetic disorder that is caused by TAZ mutations (146), resulting in lower mitochondrial CL levels and higher MLCL levels. Patients with Barth syndrome experience cardiomyopathy, skeletal muscle weakness, neutropenia, organic aciduria and growth retardation across a spectrum of severity (139).

1.4.1.4 Quality Control: Biogenesis and Mitophagy

Mitochondrial contain their own DNA genome, mtDNA, which is evidence of their prokaryotic origin and endosymbiotic relationship with cellular organisms that evolved into eukaryotes. Mitochondria cannot be generated de novo within cells, and are maternally inherited, with some exceptions (147). Multiple cellular mechanisms exist to perform mitochondrial quality control, which is essential to maintain a population of functional mitochondria to support homeostasis. The major processes of mitochondrial quality control are mitochondrial biogenesis (fusion and fission) and mitophagy

(mitochondrial autophagy).

Mitochondrial biogenesis is the expansion of mitochondrial mass within a cell’s established network of mitochondria. Increasing mitochondrial mass often occurs when cells experience heightened energy demand, although mitochondrial biogenesis is itself

an energy-consuming process. Mitochondrial biogenesis is coordinated by several transcription factors, both nuclear and mitochondrial. Peroxisome proliferator-activated receptor gamma coactivator 1a (PGC-1a) is a nuclear transcription factor that is regarded as the master regulator of mitochondrial biogenesis. It activates nuclear respiratory factor 1 (NRF1) and nuclear factor erythroid 2 related factor 2 (NRF2) (148), which regulate the expression of multiple nuclear-encoded mitochondrial proteins, including mitochondrial transcription factor A (TFAM) (149). Upon translation, TFAM is transported to mitochondria, where it acts as a transcription factor of mitochondrial proteins encoded by mtDNA and additionally coordinates mtDNA replication (150). In experimental models, increased PGC-1a expression is associated with increased mitochondrial mass, while decreased PGC-1a expression is associated with depressed mitochondrial function (149).

Mitochondrial fusion is the joining of two mitochondria to exchange materials, proteins and mtDNA. This diminishes damage to any single mitochondrion (150, 151).

To complete mitochondrial fusion, both the OMM and IMM must fuse between the two organelles. OMM tethering and fusion is mediated by the mitofusins (MFNs), dynamin superfamily transmembrane GTPases (152). Mammalian cells contain two MFN isoforms, MFN1 and MFN2. MFN1 has the most activity in mitochondrial fusion, while

MFN2 is involved in MAM tethering and mitochondrial calcium uptake (150). MFN dimerization drives OMM fusion. IMM fusion is also coordinated by a dynamin superfamily transmembrane GTPase, OPA1, which is embedded in the IMM. Proteolytic

cleavage of OPA1 into a short, non-membrane-bound form, S-OPA1 leads to S-OPA1 complexing with membrane-bound OPA1 to mediate IMM fusion (151).

Mitochondrial fission is the opposite of fusion: the division of a single mitochondrion into two organelles. This process facilitates the transmission of mitochondria to daughter cells during mitosis and sequesters damaged mitochondria for degradation. Mitochondrial fusion and fission together regulate mitochondrial morphology, which ranges from long, tubular networks to smaller, spherical mitochondria, depending on cell type and energetic needs (151). Dynamin-related protein 1 (DRP1), a cytosolic dynamin-like GTPase, orchestrates mitochondrial fission.

DRP1 is recruited to mitochondrial fission sites by OMM DRP1 receptors, including mitochondrial fission factor (MFF), fission mitochondria 1 (FIS1), and mitochondrial elongation factor (MIEF) 1 and 2. DRP1 polymerizes at these sites to form a ring-like protein complex, which leads to constriction and scission of the mitochondrion (150,

151).

The final element of mitochondrial quality control is the removal of damaged mitochondria by mitophagy, a form of selective autophagy. Multiple proteins have been identified to act as mitophagy receptors, triggering this process [reviewed in (149, 153)].

Phosphate and tensin homologue, induced putative kinase 1 (PINK1) is a serine/threonine kinase that is degraded in healthy, functional mitochondria. However, when mitochondria are damaged and depolarized PINK1 accumulates in the OMM, is autophosphorylated, phosphorylates ubiquitinated OMM proteins, and activates Parkin, an E3 ubiquitin ligase. This results in further ubiquitination and phosphorylation of OMM

proteins, culminating in the recruitment of microtubule-associated protein 1 light chain 3

(LC3), initiating the canonical autophagic pathway to target the damaged mitochondrion.

Other proteins can also recruit LC3 to mitochondria to initiate mitophagy. These include

BCL2 interacting protein 3 (BNIP3), NIX, and FUN14 domain containing 1 (FUNDC1)

(149, 150). Externalization of CL moieties on the OMM can also recruit LC3 to damaged mitochondria.

Mitochondrial Function

1.4.2.1 Mitochondrial Energy Production

The canonical role of mitochondria is as the “powerhouse of the cell”- the organelle responsible for generating energy to support cellular function. Energy production begins with the catabolism of glucose (glycolysis) or fatty acids (beta oxidation) to generate metabolites that enter the TCA cycle, which in turn provides essential electron carriers to the ETC, which generates cellular ATP via oxidative phosphorylation. Figure 1.8 shows the relationships between each of these metabolic processes.

Glycolysis occurs in the cytoplasm of cells. This process consists of a series of enzymatic reactions that break down stored glucose, a six-carbon sugar, into two three- carbon sugars, which are converted to pyruvate molecules. The catabolism of one glucose molecule via glycolysis results in two molecules of pyruvate and the generation of four molecules of ATP. Under aerobic conditions, pyruvate produced by glycolysis is transferred to the mitochondrial matrix where it is converted to acetyl-CoA by pyruvate dehydrogenase complex and enters the TCA cycle. Under anaerobic conditions (in the

absence of oxygen), pyruvate is converted to lactate in the cytosol and excreted from the cell. This reaction, catalyzed by lactate dehydrogenase, oxidizes NADH to NAD+ for use in glycolysis.

Beta oxidation occurs following transport of fatty acids into the mitochondrial matrix by carnitine translocase. Once within the matrix, fatty acid chains are cleaved into two-carbon units and converted to acetyl-CoA for the TCA cycle. The reactions required to generate acetyl-CoA from fatty acids are catalyzed by the mitochondrial trifunctional protein, which is embedded in the IMM [reviewed in (154)]. Acetyl-CoA, generated from a number of sources including glycolysis and beta oxidation, is converted to citrate at the beginning of the TCA cycle. Citrate then undergoes a series of metabolic conversions that generate NADH or FADH2 as byproducts. The final reaction of the TCA cycle generates oxaloacetate, which can be converted back into citrate to begin the cycle again. The TCA cycle is a central, highly-regulated pathway in cellular energy production, and its role in coordinating bioenergetic processes is a growing area of research across disease states [reviewed in (155)].

The final component of aerobic energy production is the ETC, which is composed of a series of enzyme complexes (I-V) embedded in the IMM. These complexes use oxidation-reduction reactions featuring the electron carriers NADH and FADH2 to pump protons from the mitochondrial matrix into the intermembrane space. Electrons from carriers are transported down the chain of enzymes and transferred to oxygen, which is reduced to water. Meanwhile, the proton gradient generated by this process drives the phosphorylation of ADP to ATP via ATP synthase (Complex IV), referred to as oxidative

phosphorylation. Protons which had been pumped out of the matrix flow down the gradient and return to the matrix via the F0 subunit of ATP synthase, which rotates and facilitates ATP synthesis by the F1 subunit [reviewed in (156)]. A diagram illustrating the electron transport chain complexes and the movement of both protons and electrons through the chain is shown in Figure 1.9.

Complex I, also known as NADH-ubiquinone oxidoreductase, transfers two electrons from NADH to ubiquinone (also known as coenzyme Q). This catalyzed reaction is coupled to the movement of four protons across the IMM. Complex I is the largest enzyme complex in the respiratory chain with 44 subunits, seven of which are encoded by mtDNA genes. Complex I is a major source of mitochondrial ROS, and the point at which NADH produced by the TCA cycle enters the ETC (157). Complex II, succinate dehydrogenase (SDH), or succinate-quinone oxidoreductase is the only enzyme complex involved in both the electron transport chain and the TCA cycle. It consists of four nuclear-encoded subunits, SDHA, B, C, and D. SDHC and SDHD are embedded in the IMM, and SDHA and SDHB protrude into the mitochondrial matrix. As part of the TCA cycle, SDH converts succinate to fumarate. This reaction is coupled with the reduction of ubiquinone. Complex II is the only ETC complex that does not pump protons across the IMM. It is also the only complex that is entirely encoded by nuclear genes (157-159). Complex III, also known as cytochrome bc1 complex or ubiquinol- cytochrome c oxidoreductase, transfers electrons from ubiquinol to cytochrome c in a process known as the ubiquinone (Q) cycle (160). This reaction is coupled with the movement of protons across the IMM. Complex III has 11 subunits, one of which is encoded by mtDNA (157). The final redox reaction of the respiratory chain occurs at

Complex IV, cytochrome c oxidase (COX). Here, electrons from cytochrome c are transferred to oxygen to make water. This reaction involves multiple co-factors, including copper ions and heme prosthetic groups. Complex IV is the only ETC complex that incorporates tissue-specific nuclear-encoded subunits that regulate complex assembly and respiratory capacity in each organ (157).

ATP synthase, F1Fo ATPase, or Complex V converts ADP and Pi to ATP in the mitochondrial matrix. ATP synthase is composed of two major subunits, F1 in the mitochondrial matrix and Fo attached to the IMM. F1 subunits δ, ε, and γ compose the complex’s central stalk. 8 copies of Fo subunit c make up a rotor embedded in the IMM below the central stalk. Fo subunits b, d, F6 and the oligomycin sensitivity-conferring protein (OSCP) form a peripheral stalk (161). This stalk keeps the F1 α and β subunits, which alternate in a ring around the central stalk, from rotating (156). The ring consists of

3 α and 3 β subunits, with the catalytic site for ATP synthesis located at the interface of each pair of subunits (161). ATP synthase is a tiny biological nanomotor, powered by the proton motive force, as described by the chemiosmotic theory (162). In this mechanism, protons sequestered in the IMS by complexes I-IV flow through the pore in Fo, driving the rotation of the Fo subunit c rotor and the connected F1 central stalk. 360° of stalk rotation forces the β subunits surrounding the stalk through a series three of conformational changes that catalyze ATP synthesis. This process is known as rotational catalysis or the binding-change mechanism (156, 161). The three conformations facilitate 1) ADP and Pi binding, 2) ATP formation and, 3) ATP release (161).

1.4.2.2 Mitochondrial Signaling

In addition to energy production, mitochondria also play an important role in cellular signaling. Calcium (Ca2+) signaling is a major cellular mechanism for perpetuating the signals of extracellular messengers. The ER is primarily responsible for regulating intracellular Ca2+ concentration, but the ability of mitochondria to take up and store Ca2+ has long been recognized (163). Mitochondrial Ca2+ uptake can buffer cytosolic Ca2+ signaling, and additionally regulate mitochondrial activity and viability.

Ca2+ concentration regulates the activity of multiple enzymes involved in mitochondrial

ATP production. The accumulation of Ca2+ in mitochondria results in an increase in mitochondrial energy production (164, 165). However, when mitochondrial Ca2+ uptake results in Ca2+ overload, it can launch cells into apoptosis. Ca2+ overload triggers formation of the mitochondrial permeability transition pore (mPTP), which leads to mitochondrial swelling and cytochrome c release into the cytosol, where it activates intrinsic apoptotic signaling pathways (164, 166).

While the role of mitochondrial in modulating cellular Ca2+ levels is well established, the role of mitochondria as a major regulator of immune system signaling has only recently been explored. Mitochondria are now known to coordinate a number of events during the innate immune response by serving as a signaling platform, as well as by releasing mitochondrial damage-associated molecular patterns (mtDAMPS) to trigger an immune reaction. Multiple families of PRRs signal through mitochondria following activation by PAMPs. The best-defined interaction is that of RLRs with MAVS. RLRs, including RIG-I, MDA5 and LGP2 detect viral RNA in the cytosol. Once activated by binding viral RNA, RLRs bind to MAVS, which is tethered to the cytosolic side of the

OMM. RLR binding activates MAVS, and triggers a prion-like MAVS polymerization that results in complexing with Tank binding kinase 1 (TBK1) and IKB kinase-e (IKKe) (167).

This complex activates transcription factors IRF3, IRF7 and NFkB, which in turn increase transcription of type I IFNs, pro-inflammatory cytokines and IFN-stimulated genes to mount an immune response to the detected pathogen (168). RLR signaling requires active oxidative phosphorylation and intact mitochondrial fusion mechanisms, and defects in these areas have been shown to lead to deficiencies in innate immune responses (169, 170).

Other PRRs also interact with mitochondrial proteins to complete their signaling cascades. Activated TLRs signal via tumor necrosis factor receptor-associated factor 6

(TRAF6), which translocates to mitochondria and ubiquitinates ECSIT, a mitochondrial protein associated with Complex I, resulting in increased mROS production, which aids in bacterial killing (168, 171). TLR7, which is activated by viral nucleic acids, stimulates

OMM ubiquitin ligase MARCH5, resulting in TANK ubiquitination. This prevents TANK inhibition of TRAF6 and enhances TLR7 signaling (172). A subset of NLRs, cytosolic

PRRs, also have mitochondrial interactions. NLR activation leads to the formation of a protein complex called the inflammasome, which activates caspase-1, IL-1b and IL-18, initiating pro-inflammatory and pro-apoptotic signaling. Following activation, NLRP3 co- localizes with mitochondria, which likely serve as a scaffold for inflammasome assembly.

Several OMM entities have been identified that interact with the NLRP3 inflammasome, including MAVS (173, 174) and cardiolipin (168, 175).

In addition to mediating response to cytosolic and endosomal PAMPs that are detected by PRRs, mitochondria also produce components that act as mtDAMPS. mtDNA release from damaged mitochondria contributes to NLRP3 inflammasome activation (176, 177). Other mtDAMPs that contribute to pro-inflammatory processes include ATP, succinate, cardiolipin and formyl peptides [reviewed in (178)]. The interactions of mitochondria with various immune signaling mechanisms are concisely illustrated in Figure 1.10.

1.4.2.3 Mitochondrial Apoptosis

Through their generation of energy, mitochondria power cell survival. However, mitochondria also play a major role in cell death. Indeed, intrinsic apoptosis, programmed cell death induced via internal signals, is also referred to as mitochondrial apoptosis. This process occurs in four general stages, beginning with activation of BCL-

2-associated X protein (BAX) and BLC-2 homologous antagonist/killer (BAK), members of the B cell lymphoma 2 (BCL-2) family of apoptosis regulatory proteins. Once activated, BAX and BAK mediate mitochondrial outer membrane permeability (MOMP), resulting in the release of soluble proteins from the intermembrane space, and eventually cell death.

Following an apoptotic stress, such as DNA damage, a subset of BCL-2 proteins known as BH3-only proteins are activated, and in turn activate BAX and BAK. Once activated, BAX and BAK localize to mitochondria and dimerize, eventually forming higher-order oligomers that are necessary for forming and stabilizing lipidic pores in the

OMM to induce MOMP (179, 180). In an apoptotic cell, all mitochondria will undergo

MOMP within a ten-minute window (181). Due to the intensity of this response, MOMP is considered the committed step of apoptosis. Following MOMP, cytochrome c is released from mitochondria. Once in the cytosol it binds an adaptor, apoptotic peptidase activating factor 1 (APAF), forming the apoptosome, which binds and activates caspase

9, the initiator caspase. Caspase 9 then cleaves and activates caspases 3 and 7, the executioner caspases. At this point, the intrinsic (mitochondrial) and extrinsic apoptotic pathways merge. Caspases 3 and 7 cleave many cellular components, leading to cell death (180). Although apoptosis is considered an anti-inflammatory process, MOMP is an inflammatory event, and contributes to several pro-inflammatory signaling pathways

[reviewed in (180)].

Mitochondria in Viral Infection

As obligate intracellular parasites, viruses are dependent on host cell resources to meet the demands of their replication cycle. This includes production of virion components, genome replication, and avoidance of anti-viral responses; all of which are energy consuming processes. Given the role of mitochondria as the major energy producer in a cell and a hub for anti-viral and apoptotic signaling; it is not surprising that many viruses interact with mitochondria during infection, either directly or indirectly.

Evidence of mitochondrial manipulation during infection has been found for diverse viruses across cell types and species [reviewed in (182-185)]. In the interest of providing relevant information, this review will focus on viruses that target the respiratory tract, with only a brief exploration of important findings in other viruses and organ systems.

One way in which viruses interact with mitochondria is to enhance viral genome replication and protein synthesis for the production of progeny virions. Acanthamoeba polyphaga mimivirus (APMV), is a giant double-stranded DNA virus that infects amoebas, but can also cause pneumonia in humans (186, 187). APMV encodes a mitochondrial transport protein called viral mitochondrial carrier 1 (VMC1). This protein removes nucleotide triphosphates, primarily dATP and dTTP, from the mitochondrial nucleotide pool to enhance replication of its own genome (182, 187). Another DNA virus,

Epstein-Barr virus (EBV), which causes infectious mononucleosis, encodes two proteins,

Zebra (BZLF1) and immediate early Zta protein, which target mitochondrial single- stranded DNA binding protein (mtSSB). BZLF1 and immediate early Zta redirect mtSSB from the mitochondria to the nucleus to support viral genome replication (188, 189).

In addition to co-opting mitochondrial components for genome replication, viruses also increase cellular energy production to support their replication cycles. Hepatitis C virus (HCV, positive-sense ssRNA) and human cytomegalovirus (HCMV, DNA) both increase mitochondrial Ca2+ concentration to increase oxidative phosphorylation and

ATP production (182, 190-192). Viruses can also target other areas of cell metabolism to increase energy capacity. Dengue virus (DENV, positive-sense ssRNA) activates AMP- activated protein kinase (AMPK) and suppresses mechanistic target of rapamycin

(mTOR), to increase b-oxidation (193, 194). HCV has also been reported to increase b- oxidation via AMPK activation (195). Conversely, RSV (negative-sense ssRNA) reduces oxidative phosphorylation, increases mROS production and orchestrates a shift to glycolysis to support viral replication in HEK293T (human embryonic kidney) and A549

(alveolar adenocarcinoma) cells (196, 197). In BALB/c mice infected with RSV,

treatment with MitoQ, a mitochondrial ROS scavenger, significantly reduced RSV viral titers in the lungs and attenuated the inflammatory response to the virus (197).

Viruses also target mitochondria to interfere with antiviral signaling. RIG-I and

MDA5 detect cytosolic viral RNA and signal through MAVS to induce antiviral gene expression. Some viruses target MAVS directly. IAV encodes PB1-F2, which is synthesized from a +1 frameshift in the ORF of PB1, a component of the viral polymerase (198). PB1-F2 targets MAVS to inhibit type I IFN induction in virus-infected cells, although the mechanism of MAVS inhibition is unclear (199). PB1-F2 can be transported into mitochondria by translocase of OMM 40 (TOM40), where it reduces mitochondrial membrane potential and induces mitochondrial fragmentation, both of which may contribute to MAVS signaling suppression (200). Interestingly, the increase in pathogenicity of the Spanish flu (H1N1) and the avian flu (H5N1) pandemic viruses is often attributed to a single mutation in PB1-F2 (201), underscoring the importance of mitochondria in the pathogenesis of viral disease. A second IAV accessory protein has also been identified that can inhibit MAVS signaling. PB2-S1 is a splice-variant encoded by PB2 with deletion of approximately 400 nucleotides (202). There are conflicting reports as to whether PB2-S1 actually binds MAVS directly, or if it destabilizes MAVS by interfering with mitochondrial integrity, but either way PB2-S1 is able to interfere with type I IFN induction downstream of mitochondria (202, 203)

Another respiratory virus capable of causing severe respiratory disease, SARS-

CoV-1 (positive-sense ssRNA), also encodes a gene that targets MAVS. Open reading frame 9b (ORF-9B) degrades MAVS, TRAF3, and TRAF6, to suppress the IFN response

(184, 204). Similarly, HCV protein NS3/4 cleaves MAVS from the OMM (182, 205).

Other viruses attenuate mitochondrial antiviral signaling by inducing mitophagy, which increases flux of mitochondrial membranes and therefore decreases the stability of OMM proteins, including MAVS [reviewed in (206)]. Viruses that employ this strategy include human parainfluenza virus type 3 (HPIV3, negative-sense ssRNA), RSV, and Measles virus (MeV, negative-sense ssRNA) (184).

A third area for virus-mitochondria interactions concerns the regulation of apoptosis. The relationship between viral replication and apoptosis is complex. Whether viruses induce or delay cell death varies depending on timing within their replication cycle, and their need to avoid immune responses, induce latency, release progeny virions, and many other factors. Whatever the intent, viruses target mitochondria to modulate apoptosis because mitochondria serve as a cell death signaling hub. In addition to using mitophagy to attenuate IFN induction, a number of viruses also use mitophagy to prevent apoptosis by blocking cytochrome c release. Hepatitis B virus

(HBV, dsDNA), HCV, Venezuelan equine encephalitis virus (VEEV, positive-sense ssRNA), classical swine fever virus (CSFV, positive-sense ssRNA), porcine reproductive and respiratory syndrome virus (PRRSV, positive-sense ssRNA), Newcastle disease virus (NDV, negative-sense ssRNA), and transmissible gastroenteritis virus (TGEV, positive-sense ssRNA) all employ such strategies to prevent apoptosis (206). Similarly, herpes simplex virus type 1 (HSV-1, dsDNA) and vaccinia virus (dsDNA) encode anti- apoptotic viral proteins that prevent cytochrome c release from mitochondria (207, 208).

Other mitochondrial-targeted strategies induce apoptosis. Human papillomavirus

(HPV, dsDNA) encodes a protein that binds mitochondria and causes their detachment from microtubules, leading to a reduction in mitochondrial membrane potential and apoptosis induction (209). Human immunodeficiency virus (HIV, positive-sense ssRNA) encodes several proteins that promote cell death. Vpr disrupts Mfn2-ER interactions, reducing OMM integrity, and binds IMM adenine nucleotide translocase (ANT) to generate a pro-apoptotic pore (210, 211). Gp120 complexes with CXCR4 in CD4+ T cells and induces MOMP and cytochrome c release (212). Another retrovirus, human T- cell leukemia virus (HTLV) causes cell death by inducing a potassium influx into mitochondria that leads to fragmentation and apoptosis (184, 213).

Multiple IAV proteins have been identified that induce or prevent apoptosis, several of which interact with mitochondria [reviewed in (214)]. In addition to its anti-

MAVS role, PB1-F2 can also induce MOMP and cytochrome c release triggering intrinsic apoptosis (198, 215). A subset of NS1 has also been reported to localize to mitochondria

(216), and this is associated with induction of intrinsic apoptosis in highly pathogenic avian IAV H5N1 (217).

Mitochondria in Lung Disease

Over the past decade, mitochondria have emerged as a major contributor to the progression of multiple pulmonary diseases, including chronic obstructive pulmonary disorder (COPD), pulmonary fibrosis (PF), and asthma. More recently, examination of the role of mitochondria in ARDS has begun. For a better understanding of the role of mitochondria in lung function, we will review the contributions of mitochondria to each

disease state, beginning with chronic disease, and then discuss the role of mitochondrial dysfunction in ARDS.

COPD is a complex lung condition characterized by chronic inflammation, emphysema and airway obstruction that leads to a progressive and substantial decline in lung function. Cigarette smoking is a major cause of COPD, but chemical and pollution exposures contribute as well, as do underlying genetic factors. Lung epithelial cells are thought to drive the inflammatory processes that lead to COPD (218). Epithelial cells in lungs from COPD and emphysema patients exhibit abnormal mitochondrial morphology, including swollen, fragmented mitochondria with loss of cristae and abnormal branching patterns (219-221). Three months exposure of mice to cigarette smoke, an in vivo model of COPD, also led to mitochondrial dysfunction, including abnormal morphology, reduced mtDNA and ATP levels, reduced rates of oxygen consumption and ETC activity, loss of mitochondrial membrane potential and increased mitochondrial ROS (mtROS)

[reviewed in (218)]. In a low-dose cigarette smoke exposure model, mitochondrial hyperfusion and increased metabolic activity were reported in primary murine ATII cells

(222), suggesting a period of mitochondrial adaptation, known as mitohormesis, before dysfunction develops (218).

As previously discussed, cells rely on mitochondrial biogenesis and mitophagy to repair and expand mitochondrial capacity. Expression of PGC1-a, the master regulator of mitochondrial biogenesis is increased in lungs from patients with mild COPD, but returns to control levels as disease severity increases (223), supporting the idea of transient mitohormesis. Increased PGC1-a transcription in COPD and cigarette smoke

exposure models has also been reported (220, 224). Transcription and expression of

PINK1, a master regulator of mitophagy, are increased in lung epithelial cells from

COPD patients as well (220, 225). Similarly, DRP1, a mitochondrial fission protein, was increasingly phosphorylated and activated in lungs of mice exposed to cigarette smoke and alveolar epithelial cells obtained from human smokers (221, 225). Overall, an upregulation of mitochondrial quality control, at least transiently, is associated with

COPD pathogenesis.

COPD is inflammatory disease, and multiple mitochondria components are able to activate pro-inflammatory pathways. COPD is associated with increased mtROS generation, and mtROS can activate NF-kB, HIF-1a and the inflammasome (168). mtDAMPs also activate inflammatory responses (168, 170). Cigarette smoke exposure is associate with increased serum and bronchoalveolar lavage fluid (BALF) mtDNA in mice (226, 227), and patients with COPD have increased extracellular ATP (228, 229).

Both mtDNA and ATP are potent mtDAMPs that may elicit a pro-inflammatory response, contributing to COPD pathogenesis. Mitochondria are also centers of iron homeostasis, and recent research suggests that iron overload may contribute to mitochondrial dysfunction observed in COPD (230, 231).

Idiopathic pulmonary fibrosis (IPF) is another chronic lung disease in which mitochondria may play a role. IPF is considered a disease of aging and is characterized by abnormal scarring of the lung parenchyma that causes a progressive decline in lung function. A number of indicators of mitochondrial dysfunction have been observed in alveolar epithelial cells in lungs of IPF patients (232). These include abnormal

mitochondrial morphology, and reduced expression of PINK1, suggesting a reduction in mitophagy (233). The role of PINK1 downregulation as a factor in pulmonary fibrosis development has been confirmed in multiple mouse models (233-235). Mitochondrial quality control deficiencies in alveolar epithelial cells may play a broader role in IPF, as mitochondrial fusion is also dysregulated. Cells from IPF patients over-express MFN2, and disruption of mitochondrial fusion in a mouse model of IPF drives fibrosis (236).

Fibroblasts, which are aberrantly activated to a pro-fibrotic phenotype in IPF, undergo metabolic reprogramming to aerobic glycolysis, which may involve changes in mitochondrial energy production (237, 238). Lungs from IPF patients exhibited elevated

ROS, and signs of reduced ETC activity and mtDNA damage (239, 240). Also of note,

TGF-b, a canonical driver of fibrotic remodeling in the lung, also plays a role in modulating mitochondrial quality control mechanisms, and is activated by elevated mtROS levels (232, 239, 241).

In contrast to COPD and IPF, which are associated with aging, asthma often develops in childhood. Asthma is a disease of airway hyperresponsiveness, characterized by episodes of bronchoconstriction, increased mucus secretion, coughing, wheezing and shortness of breath, often triggered by an allergen or other irritant. Both airway smooth muscle (ASM) and airway epithelial cells are involved in asthma pathophysiology. Both cell types exhibit abnormal mitochondrial morphology and elevated mitochondrial Ca2+ levels in samples from asthma patients as well as allergic asthma mouse models (242, 243). Asthmatic ASM cells exhibit increased mitochondrial mass and mitochondrial respiration, possibly to respond to increased energy demand for cell growth and proliferation during airway remodeling (242). Increased mitochondrial

biogenesis in asthma is accompanied by an increase in DRP1 expression and a concomitant decrease in MFN2 expression, suggesting upregulation of mitochondrial fission and a reduction in fused networks of mitochondria (242, 244). Overall, asthma is associated with changes in mitochondria that expand energy production capacity.

Mitochondrial dysfunction has also been observed in acute respiratory failure and illness that leads to ARDS, although much of this data has been generated in pre-clinical models. In rabbits with LPS-induced ARDS, lung ATP content was depleted, as were the levels of bioenergetic metabolites, such as ADP and NAD+ (245). In line with this data, a mouse model of LPS-induced ARDS showed reduced lung ATP and mitochondrial respiration (246, 247). Chemically-induced lung injury depolarized alveolar endothelial cell mitochondria in mice, resulting in endothelial barrier failure (248) Sepsis, a major cause of ARDS, increases mtROS and damages the ETC in multiple tissues, including cardiac muscle, skeletal muscle, and peripheral blood monocytes, resulting in impaired oxidative phosphorylation and ATP production (249, 250). A study in patients with septic shock found that poor mitochondrial function was correlated with increased mortality risk

(250). Multiple in vivo models of sterile inflammation have linked extracellular mtDNA release to acute lung injury induced by acid aspiration, surgical trauma, and acute drug toxicity (251, 252). Research has also indicated that circulating mtDNA is a valid biomarker for sepsis [reviewed in (253)]. A final line of evidence that underscores the importance of mitochondrial dysfunction in ARDS is the capacity of mitochondrial transfer between cells to protect against acute lung injury. In mouse models of ARDS induced by LPS or E. coli instillation, subsequent instillation of bone marrow-derived mesenchymal stromal cells (MSCs) attenuated clinical signs of ARDS as a result of

mitochondrial transfer from MSCs to alveolar macrophages and epithelial cells (254,

255).

1.5 Figures

Taubenberger JK and Kash JC. Cell Host Microbe. 2010.

Figure 1.1: Influenza A virus virion composition.

Knipe DM, Howley P. Fields Virology. Philadelphia: Wolters Kluwer; 2013.

Figure 1.2: Multiple RNA products are derived from a single IAV vRNA.

Huppert LA and Matthay MA. Front Immunol. 2017.

Figure 1.3: Vectorial ion transport and alveolar fluid clearance.

Agassandian M and Mallampalli RK. Biochim Biophys Acta. 2013.

Figure 1.4: Synthesis, secretion and recycling of pulmonary surfactant.

Mejia EM and Hatch GM. J Bioenerg Biomembr. 2015.

Figure 1.5: Mitochondrial structure and phospholipid composition

[Orange = PC, yellow = PE, white = CL].

Figure 1.6: Kennedy pathway de novo synthesis of phosphatidylcholine.

Mejia EM et al. Chem Phys Lipids. 2014.

Figure 1.7: CDP-DAG pathway de novo synthesis and remodeling of cardiolipin.

Nsiah-Sefaa A and McKenzie M. Biosci Rep. 2016.

Figure 1.8: An overview of cellular energy metabolism as it relates to mitochondria.

Rotig A and Munnich A. J Am Soc Nephrol. 2003

Figure 1.9: Movement of protons and electrons through the electron transport chain complexes.

Banoth B and Cassel SL. Transl Res. 2018

Figure 1.10: Mitochondria play a role in multiple immune signaling pathways.

1.6 Tables

Adapted from The ARDS Definition Task Force. JAMA. 2012.

Table 1.1: Berlin definition of ARDS.

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Chapter 2. Acutely Lethal Influenza Infection Alters ATII Cell Mitochondrial Structure and Function

2.1 Abstract

Influenza A virus (IAV) is a zoonotic respiratory virus that causes severe disease and substantial mortality during annual seasonal influenza epidemics. Although many cases of IAV infection are mild, those that progress to severe disease often involve IAV replication in alveolar type II (ATII) cells in the lower respiratory tract. ATII cells are essential for maintaining normal lung function, and development of acute respiratory distress syndrome (ARDS) secondary to severe influenza is associated with defects in

ATII cell function. ARDS is a highly lethal disease characterized by hypoxemia, pulmonary edema, reduced lung compliance and decreased gas exchange. Influenza is the leading viral cause of ARDS in humans. Our previous work has shown that reduction in lung compliance in a mouse model of ARDS is correlated with defects in ATII cell pulmonary surfactant production. Phosphatidylcholine (PC) and phosphatidylethanolamine (PE) are the major components of pulmonary surfactant, which facilitates lung compliance by reducing alveolar surface tension. PC and PE synthesis are disrupted in ATII cells following in IAV infection, resulting in significantly lower levels of these phospholipids. PC and PE are also the major phospholipids in mitochondrial membranes. Mitochondria produce the majority of the energy needed for ATII cell activities by oxidative phosphorylation, which requires intact mitochondrial membranes to generate proton motive force and drive ATP synthesis. We therefore hypothesized that reduction in PC and PE availability following IAV infection would impact ATII cell mitochondrial structure and function. We found that ATII cells from IAV-infected mice

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have abnormal mitochondrial ultrastructure, increased frequency of mitochondrial reactive oxygen species (mtROS), altered cardiolipin synthesis, and up-regulated mitochondrial biogenesis. These cells exhibit mitochondrial depolarization and reduced

ATP production following infection and the development of ARDS. However, these changes are not due to the initiation of apoptosis or defects in electron transport chain enzymes. Overall these results suggest that metabolic and structural changes may underlie observed defects in mitochondrial energy production in ATII cells from IAV- infected mice. ATII cell energy deficits due to mitochondrial dysfunction may alter ATII cell function and reduce their ability to maintain alveolar homeostasis, therefore contributing to the development of ARDS following IAV infection.

2.2 Introduction

Influenza A virus (IAV) causes yearly seasonal influenza epidemics and has demonstrated pandemic potential (1). Many influenza cases consist of respiratory and systemic symptoms, such as cough, sore throat, fever, headache, and malaise that will resolve on their own in about a week’s time (2). However, influenza can progress to severe disease, which can manifest with an aberrant immune response, primary viral pneumonia, secondary bacterial pneumonia, cardiac complications, neurological issues and kidney failure, and requires hospitalization (3). Over 50% of severe influenza patients in the intensive care unit (ICU) will develop acute respiratory distress syndrome

(ARDS) (3). ARDS is a type of acute respiratory failure that is caused by diverse insults, including viral infections of the respiratory tract, and characterized by substantial morbidity and mortality. ARDS can only be treated with supportive care, primarily

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mechanical ventilation (4), and 40% of patients diagnosed with ARDS will die while hospitalized for treatment (5).

In the United States alone, our healthcare system treats about 200,000 ARDS patients each year, resulting in about 75,000 deaths (4). The current coronavirus disease 2019 (COVID-19) pandemic has the potential to significantly increase the burden of this syndrome, as early estimates suggest that up to 40% of hospitalized

COVID-19 cases will progress to ARDS (6). It is therefore evident that substantial research effort should be put towards developing therapeutic strategies to advance

ARDS treatment beyond supportive care and reduce the toll of this deadly diagnosis.

To address this therapeutic gap, we utilize a mouse model of IAV-induced ARDS that recapitulates the clinical signs of ARDS observed in humans: acute hypoxemia, pulmonary edema, decreased lung compliance, and reduced gas exchange, indicated by a PaO2:FiO2 ratio of below 200 mmHg (7-9). Lung compliance is normally maintained by pulmonary surfactant, a complex mixture of proteins and lipids that is secreted into the alveolar space by alveolar type II epithelial (ATII) cells. There are four surfactant proteins

(SP): SP-A, SP-B, SP-C, and SP-D, which are expressed exclusively by ATII cells in the lung (10). Surfactant proteins make up 10% of pulmonary surfactant, while lipids make up 90%. The majority of the lipid component of pulmonary surfactant is phospholipids, the most important of which are phosphatidylcholine (PC) and phosphatidylethanolamine

(PE). Like surfactant proteins, surfactant phospholipids are synthesized in ATII cells

(11). Defects in surfactant production and secretion can lead to respiratory failure (12).

We and others have reported that IAV infection reduces surfactant protein synthesis in murine ATII cells (13, 14), and we have subsequently shown that production

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of surfactant phospholipids in ATII cells is also affected by IAV infection (15). Our metabolomic analysis of ATII cells from IAV-infected mice identified both PC and PE levels as being significantly reduced at 6 days post-inoculation (dpi). In addition, critical intermediate metabolites for de novo PC and PE synthesis via the Kennedy pathway

(16) were also present at significantly lower levels, suggesting defects in PC and PE production in ATII cells from IAV-infected mice (15).

Not only are PC and PE essential components of pulmonary surfactant, they are also the major phospholipids in mitochondrial membranes (17). Mitochondria, the powerhouse of the cell, are composed of two phospholipid bilayers, the outer mitochondrial membrane (OMM) and the inner mitochondrial membrane (IMM). The

OMM coordinates protein trafficking and signaling between the mitochondria and the rest of the cell, while the IMM is primarily concerned with energy production- oxidative phosphorylation to produce adenosine triphosphate (ATP) via the electron transport chain (ETC). These two membranes are critical for cellular and organismal survival, as the mitochondria produce the majority of the energy required for cellular function. The

IMM surrounds the mitochondrial matrix, the innermost compartment of this organelle, where metabolic reactions to support ATP synthesis occur, and where mtDNA is stored.

A number of important metabolic reactions that support ATP production take place in the matrix, including those in the tricarboxylic acid (TCA) cycle and fatty acid oxidation (18,

19). The area between the IMM and OMM is the intermembrane space, where protons used to drive ATP synthase are stored during oxidative phosphorylation. The OMM contains 54% PC and 29% PE, while the IMM contains 40% PC and 34% PE, as well as

18% cardiolipin, a phospholipid specific to mitochondrial membranes (17, 20, 21).

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Considering the importance of PC and PE for forming mitochondrial membranes, we hypothesized that observed changes in ATII cell phospholipid levels following IAV infection would alter mitochondrial structure and function, as mitochondrial membranes are predominantly composed of phospholipids with specific physical properties that support mitochondrial physiology and intact membrane structure is critical for ETC ATP synthesis. Changes in mitochondrial structure and/or function have been previously observed in pulmonary epithelial cells in emphysema, chronic obstructive pulmonary disease (COPD) (22-24), idiopathic pulmonary fibrosis (IPF) (25), and asthma (26-28).

To identify for the first time changes in ATII cell mitochondria in vivo during

ARDS, we examined aspects of mitochondrial ultrastructure, morphology, and physiology in ATII cells isolated from mice infected with IAV. To assess changes in ATII cell mitochondrial function induced by IAV infection, we measured multiple parameters of mitochondrial energy production by the electron transport chain (ETC). We found that mitochondrial structure is indeed altered under these pathologic conditions, and that IAV infection does indeed disrupt mitochondrial energy production, although expression of

ETC enzymes does not change significantly. Overall, reduction in ATP production by mitochondria may contribute to reported defects in ATII cell function during IAV-induced

ARDS, as the multifunctional nature of these cells results in substantial ATP demand

(29, 30).

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2.3 Materials and Methods

Animal experiments. All animal experiments complied with the NRC Guide for the Care and Use of Laboratory Animals and were approved by The Ohio State University’s

Institutional Animal Care and Use Committee.

Mouse infection. 8-12 week old female C57BL/6 mice (Charles River Laboratories,

Ashland, OH, USA) were inoculated intranasally with 10,000 plaque-forming units (pfu) of influenza A/WSN/33 (H1N1) virus in 50 microliters (μl) phosphate-buffered saline

(PBS) with 0.1% bovine serum albumin (BSA) under ketamine/xylazine anesthesia (14).

This inoculum results in 100% mortality by 8 days post-inoculation (dpi), without replication in the brain (8, 9, 31). Control mice were “mock” inoculated with 50 μl PBS with 0.1% BSA under ketamine/xylazine anesthesia to mimic the route of viral infection.

Prior to inoculation, mice were individually numbered and weighed. Mice were re- weighed every two days following infection to confirm disease progression.

ATII cell isolation. ATII cells were isolated from euthanized mice at 6 dpi using the following procedure: immediately following euthanasia by ketamine/xylazine overdose and severing of the renal artery, the thoracic cavity was carefully opened without puncturing lungs. The heart was perfused with 15-30 milliliters (mL) heparin in PBS (20 units/mL). Next, the trachea was exposed and sliced open, so that a cannula (created from an 18-gauge catheter) could be inserted. The lungs were then inflated with 1 mL of dispase II (9 mg/mL) administered via 1 mL luer slip tip syringe inserted into the cannula.

Immediately after dispase instillation, 0.6 to 0.8 mL of warm 0.1% low melting point agarose in PBS was introduced to the lungs using a second syringe, and the thoracic cavity was completely covered with ice for two minutes, until the agarose solidified into a

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“plug”. Instillation of the agarose forces the dispase into the distal airways and alveoli and keeps it from spilling back out of the lungs. Once the agarose had solidified, the lungs were excised from the thoracic cavity, taking care to avoid puncturing the now- inflated lungs, and to remove any non-lung tissue, such as the thymus, heart and trachea. The lungs were then transferred to a 15 mL conical tube containing 4 mL dispase (9 mg/mL) for digestion.

Dispase-inflated lungs were incubated on a nutating rocker at room temperature for 45 minutes. At 15, 30, 35, 40 minutes into the incubation period, tubes were shaken manually to assist in tissue break down. Following 45 minutes of incubation in dispase, 5 mL of DNAse I in DMEM (0.1 mg/mL) was added to each sample for an additional 5- minute incubation. This combination of chemical and mechanical digestion gently breaks lung tissue down into a single cell suspension with minimal cell death (<1%). Dispase II cleaves type IV collagen and fibronectin (32), leaving airway tissue intact. Following

DNAse digestion, lung samples were placed on ice, and then individually emptied into a plastic 10 cm petri dishes. Airway tissue was removed from the single cell suspension, and a 10 mL serological pipette was used to aspirate and dispense the suspension in the petri dish several times to break up any remaining clumps. The entire cell suspension was then aspirated into the serological pipette and dispensed into a 100 micron cell strainer placed into a 50 mL conical tube on ice. This process was repeated for each lung sample.

Once all lung suspensions were filtered into 50 mL conical tubes, the tubes were centrifuged at 1,000 revolutions per minute (rpm) at 4°C for 5 minutes to pellet the cells.

The supernatant was then discarded, and cells resuspended before adding 3 mL ACK

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lysing buffer for 3 minutes to lyse any red blood cells not removed by lung perfusion.

This step is especially critical for IAV-infected mice at 6 dpi, when the lungs are severely injured with substantial consolidation and difficult to perfuse. After 3 minutes, 7-10 mL of

PBS with 10% fetal bovine serum (FBS) was added to each sample to halt hemolysis reaction. After mixing, each sample was aspirated and dispensed by serological pipette

(10 mL) into a 20 micron filter in a new 50 mL conical tube on ice. 20 micron filters were created using 20 micrometer (μm) opening nylon mesh acquired from an industrial supply company. This smaller filter size will remove larger lung cells, including ATI cells

(100 μm in diameter), while ATII cells (10 μm in diameter) will pass through (33).

ATII cells were isolated from this size-filtered “crude” lung cell population using negative selection and magnetic separation. First cells were counted to determine the appropriate amount volume of selection antibodies and magnetic beads for each sample. Crude cell counts from mock-inoculated (healthy) mice ranged between 1 and

5×107 cells, while counts for IAV-infected mice ranged between 3 and 8×107 cells due to the influx of immune cells into the lungs. For healthy mice, cells were incubated with monoclonal anti-CD45, anti-CD16/32, anti-CD31, anti-TER119 and anti-CD104, each at a volume of 50 μL/107 cells. For infected mice, the volumes of anti-CD45 and anti-

CD16/32 were both increased to 70 μL/107 cells to account for the higher populations of infiltrating immune cells in these samples. Each antibody targets a different cell type that will be removed from the sample during negative selection, so that only ATII cells remain following magnetic separation. Anti-CD45 (clone 30-F11) targets hematopoietic cells and alveolar macrophages, anti-CD16/32 (clone 93) targets alveolar macrophages, anti-

CD31 (clone MEC13.3) targets endothelial cells, anti-TER119 (clone TER-119) targets

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erythroid cells, and anti-CD104 (integrin β4; clone 346-11A) targets club cells and distal lung progenitor cells (34). All antibodies were custom ordered with biotin conjugates from BioLegend. Following addition of antibodies, cell suspensions were incubated on ice for 20 minutes, mixing each sample by hand every 5 minutes. Next, 2-4 mL isolation buffer (PBS with 2% BSA and 0.01% NaN3) was added to each sample and the suspensions were centrifuged at 1,000 rpm at 4°C for 5 minutes to remove unbound antibodies. After discarding supernatant and replacing with 2-4 mL fresh isolation buffer, centrifugation was repeated to remove any remaining antibodies not bound to cells.

To prepare for magnetic separation, samples now containing lung cells bound by biotinylated antibodies were next incubated with streptavidin-conjugated magnetic beads

(Miltenyi Biotec). Beads were added to samples at a concentration of 20 μL/107 cells, based on the crude suspension cell counts obtained previously. This was followed by another 20 minute incubation on ice with mixing by hand at 5 minute intervals. Then 2-4 mL isolation buffer was added to each sample, and the suspensions were centrifuged at

1,000 rpm at 4°C for 5 minutes to remove unbound streptavidin beads. At this point, single-use magnetic columns were inserted into a magnetic separator (LS columns and

QuadroMACS Manual Separator, both Miltenyi Biotec) and washed with 3 mL isolation buffer. Following centrifugation, the supernatant was removed from samples, and cells were resuspended in 500 μL isolation buffer and transferred to columns (one sample per column). Each sample was allowed to move through column and into a clean tube until the reservoir on top of the column was almost empty, at which point 3 mL of isolation buffer was added to the reservoir. Following three such washes, the column was allowed to run until no liquid remained, at which point samples, now containing only ATII cells,

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were removed from the separator and centrifuged at 1,000 rpm at 4°C for 5 minutes to pellet cells. After discarding supernatant, cells were re-suspended in 1 mL isolation buffer and counted again. From here, isolated ATII cells were ready for use in other experiments. This procedure expands upon ATII cell isolation processes previously used in the field (14, 33-35) to deliver a population of highly pure ATII cells from an IAV- infected mouse lung, which differs significantly from a health lung. To our knowledge, this is the first protocol developed to successfully isolate ATII cells from these fragile, inflamed lungs at consistently high purity. This procedure will yield between 1 and 5×106

ATII cells per mouse.

Live cell isolation. Live ATII cells were sorted from total ATII cell populations isolated by standard protocol on an FACSAria III fluorescent cell sorter (BD Biosciences), based on Live/Dead Fixable Violet Dead Cell Stain (Life Technologies) fluorescence

(emission/excitation 416/451 nm). Live cells, which have a low fluorescent signal using this stain, were sorted into phosphate-buffered saline (PBS), which was then replaced with Qiazol (Qiagen) for cell lysis, fixation and nucleic acid extraction.

Transmission electron microscopy and image analysis. Whole lungs were perfusion- fixed with glutaraldehyde and prepared for transmission electron microscopy analysis by standard methods (36). Ultrastructure was visualized using a JEM-1400 transmission electron microscope (JEOL, Peabody, MA) linked to an Olympus SIS Veleta 2K camera

(Olympus Soft Imaging Solutions). Quantitative image analysis was performed using

ImageJ, and organelle measurements were normalized to total cell cross-section area.

Mitochondrial mass quantification. ATII cell mitochondrial mass was quantified on an

Attune Nxt Flow Cytometer (Invitrogen) using MitoTracker Red FM dye

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(excitation/emission 581/644 nm, Invitrogen). ATII cell suspensions were incubated with

250 nM MitoTracker for 30 minutes at 37°C, then the dye was removed by centrifugation at 1000 rpm for 5 minutes at 4°C. Cells were re-suspended in 1 mL FACS buffer (PBS with 2% BSA and 0.01% NaN3) and read immediately on the flow cytometer using the following parameters: forward scatter (FSC) 160V, side scatter (SSC) 345V, red laser 1

(RL1) 320V.

Mitochondrial ROS quantification. ATII cell mitochondrial ROS was quantified on an

Attune Nxt Flow Cytometer (Invitrogen) using MitoSOX Red Mitochondrial Superoxide

Indicator (510/580 nm, Life Technologies). 50 μM MitoSOX dye solution was added to cell suspension for a 10 minute incubation at 37°C. Positive control samples were generated by incubating cells with 1 μM rotenone/antimycin A (Agilent Technologies) for

10 minutes at 37°C prior to adding MitoSox probe. Following incubation, the dye was removed by centrifugation at 1000 rpm for 5 minutes at 4°C. Cells were re-suspended in

1 mL FACS buffer and read immediately on the flow cytometer using the following parameters: FSC 160V, SSC 345V, blue laser 2 (BL2) 310V.

Cardiolipin quantification. ATII cell CL content was quantified on an Attune Nxt Flow

Cytometer (Invitrogen) using nonyl acridine orange dye (NAO, 488/530 nm, Invitrogen)

(37). NAO was dissolved in 100% ethanol to make a working solution of 35 ng/mL. 500

μL NAO solution was added to each cell sample, and incubated for 10 minutes at 37°C.

Then the dye was removed by centrifugation at 1000 rpm for 5 minutes at 4°C. Cells were re-suspended in 1 mL FACS buffer and read immediately on the flow cytometer using the following parameters: FSC 160V, SSC 340V, blue laser 1 (BL1) 240V.

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Quantitative real-time PCR (qRT-PCR). RNA was extracted from ATII cells using

Qiazol and the miRNeasy Mini Kit (both Qiagen). RNA quality and quantity were assessed by spectrophotometry (NanoDrop2000). For qRT-PCR of cardiolipin synthesis pathway and mitochondrial fusion genes, total cDNA was synthesized from up to 2 micrograms (μg) of RNA using the High Capacity cDNA Reverse Transcription Kit

(Applied Biosystems). qRT-PCR assays used TaqMan primers with a FAM reporter for murine CRLS1, TAZ, MFN1 and MFN2 (Thermo Fisher), and TaqMan Universal PCR

Master Mix (Applied Biosystems). All qRT-PCR experiments were run on a 96-well

StepOnePlus Real-Time PCR System (Applied Biosystems) using the associated

StepOnePlus software. CT values were exported to Microsoft Excel and gene expression of CRLS1, TAZ, MFN1 and MFN2 was evaluated using the ΔΔCT method

(38). 18s ribosomal RNA was used as an endogenous control gene.

For the Mouse Mitochondrial Energy Metabolism PCR Array, cDNA was synthesized using the RT2 PreAMP cDNA Synthesis Kit and RT2 PreAMP Pathway

Primer Mix for Mouse Mitochondrial Energy Metabolism PCR Array (all Qiagen). PCR array plates were run using RT2 Profiler PCR Arrays with RT2 SYBR Green qPCR

Mastermix (both Qiagen). All qRT-PCR experiments were run on a 96-well StepOnePlus

Real-Time PCR System (Applied Biosystems) using the associated StepOnePlus software. CT values were exported to Microsoft Excel and gene expression patterns were evaluated using the ΔΔCT method (38). For PCR arrays, β-actin was included on the array plate and used as a housekeeping gene.

Western blotting. For relative quantification of tafazzin and PGC-1α protein expression, freshly isolated ATII cells were vortexed then frozen at -80°C in 1:10 diluted Cell Lysis

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Buffer (Cell Signaling) containing 1 μL/mL benzonase nuclease (Sigma Aldrich). After freezing, cell lysates were thawed on ice and vortexed at maximum speed in a benchtop centrifuge at 4°C for 10 minutes to remove cellular debris. Supernatants were transferred to new 1.5 mL microcentrifuge tubes and stored on ice. To determine protein concentration, BCA assays were performed according to manufacturer instructions

(Pierce BCA Protein Assay Kit) and read on a colorometric plate reader at 562 nm.

Based on BCA results, protein concentrations were calculated from the generated standard curve. To prepare samples for SDS-PAGE, 10 ug of protein was mixed with appropriate volume of 4x Bolt LDS Sample Buffer and 10x Bolt Reducing Agent (both

Life Technologies), as well as PBS to achieve a uniform loading volume across samples.

Samples were loaded into Bolt 4-12% Bis Tris Plus gels in the Invitrogen Mini Gel Tank filled with 1:20 diluted 20x Bolt MOPS SDS Running Buffer (all Life Technologies). A dual color/fluorescence molecular weight ladder (GE Healthcare) was also loaded onto the gel, which was run at 150 volts for about 1 hour, until dye front had migrated to the bottom of the gel. Protein was transferred onto an Immobilon-P PVDF membrane

(Millipore) using the Invitrogen Mini Blot Module and 1:20 diluted Bolt 20x Transfer

Buffer with 20% methanol and Bolt Antioxidant (all Life Technologies), following a standard membrane transfer protocol.

5% BSA in TBS with 0.1% Tween (TBS-T), and incubated overnight with rocking at 4°C.

The following day, the primary antibody solution was discarded, and the membrane

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washed in TBS-T three consecutive times for 10 minutes each. Membranes were then incubated on a rocker at room temperature for 1 hour with the appropriate Cy5- conjugated secondary antibody (anti-mouse or anti-goat; Thermo Fisher) diluted to

1:5000 in 5% BSA in TBS-T. Following three additional washes with TBS-T, membranes were transferred to a piece of thick filter paper and dried at 37°C for 5-10 minutes. Once dry, membranes were imaged on an Amersham Typhoon using the Cy5 670BP30 laser, which picks up the least background fluorescence on PVDF membranes. Quantification of bands was performed on .tif image files captured by the Typhoon using ImageJ software and a standard procedure. Expression of β-actin was used for normalization.

Mitochondrial membrane potential assay. ATII cell mitochondrial membrane potential was quantified on an Attune Nxt Flow Cytometer (Invitrogen) using the MitoProbe

DiIC1(5) kit (excitation/emission 633/670 nm, Invitrogen). ATII cells were incubated with

10 μM DiIC1(5) in for 30 minutes at 37°C. Positive control samples were incubated with

100 μM CCCP, a membrane potential dissipator, for 5 minutes at 37°C, prior to addition of DiIC1(5). Following incubation, the dye was removed by centrifugation at 1000 rpm for

5 minutes at 4°C. Cells were re-suspended in 1 mL PBS and read immediately on the flow cytometer using the following parameters: FSC 160V, SSC 340V, RL1 290V.

Measuring mitochondrial oxygen consumption rate (OCR). ATII cell OCR was measured using a Seahorse XFe24 Bioanalyzer and XFe24 Mito Stress Test Kit (both

Agilent). ATII cells were plated in specialized 24-well Seahorse cell culture microplates

(Agilent) at a density of 1×105 cells per well, in warm pH 7.4 DMEM with 1.0 M glucose,

100 mM pyruvate, 200 mM glutamine (assay media, all Agilent). To immobilize cells, microplates were coated with 22.4 μg/mL Cell-Tak cell and tissue adhesive (Corning) 24

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hours prior to plating. Once cells were plated, the plate was centrifuged at 300 rpm for 2 minutes with no braking to adhere cells to Cell-Tak coating. The microplate was then incubated without CO2 at 37°C for 30 minutes to equilibrate cells.

For 24 hours prior to experiment, a sensor cartridge was hydrated by incubation in Seahorse XF Calibrant solution (Agilent) without CO2 at 37°C. While cells were adhering to the microplate, the sensor cartridge was removed from the incubator, and

Mito Stress Test compounds were resuspended in assay media and loaded into cartridge injector ports at the following concentrations: 1 µM oligomycin (glycolysis inhibitor), 4 µM FCCP (proton gradient de-coupler), and 0.5 µM rotenone/antimycin A

(ETC inhibitors). The sensor cartridge was then loaded into the Seahorse analyzer for calibration. Once calibration was complete, the microplate containing ATII cells was loaded into the analyzer for data collection using Wave software (Agilent). Each experiment consisted of a series of OCR measurements, starting with 3 basal measurements, followed by injection of oligomycin into each well, mixing, 3 measurements, injection of FCCP, mixing, 3 measurements, injection of rotenone/antimycin A, 3 measurements. Once complete, readings were exported to

Microsoft Excel and the mean OCR for each sample under each condition was determined, with standard error.

Mitochondrial transition pore opening assay. ATII cell mitochondrial transition pore opening was quantified on an Attune Nxt Flow Cytometer (Invitrogen) using the

MitoProbe Transition Pore Assay kit (488/530 nm, Invitrogen). ATII cells were incubated with 10 μM calcein AM in HBSS for 15 minutes at 37°C. Mitochondrial fluorescence was measured in samples co-incubated with 10 μL CoCl2. Positive control samples were

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incubated with 10 μM calcein AM, 10 μL CoCl2, and 10 μL ionomycin . Following incubation, the dye was removed by centrifugation at 1000 rpm for 5 minutes at 4°C.

Cells were re-suspended in 1 mL HBSS and read immediately on the flow cytometer using the following parameters: FSC 160V, SSC 340V, blue laser 1 (BL1) 240V.

Statistical analysis. Descriptive statistics (mean and standard error) were calculated using Instat software (GraphPad, San Diego, CA). Gaussian data distribution was verified by the method of Kolmogorov and Smirnov. An unpaired Student’s t-test was used when comparing 2 groups. Statistical analyses of datasets containing more than 2 groups were made by ANOVA, with a post hoc Tukey-Kramer multiple comparison post- test. All data are presented as mean ± S.E.M. P<0.05 was considered statistically significant. Data undergoing statistical analysis was derived from no less than 2 separate infection groups.

2.4 Results

ATII cell mitochondrial ultrastructure is altered by IAV infection. To assess whether changes in ATII cell phospholipid synthesis would affect mitochondrial structure, we used transmission electron microscopy (TEM) to observe ultrastructural changes in ATII cell mitochondria in lungs from mock- and IAV-infected mice at 6 dpi. ATII cells were identified based on presence of characteristic lamellar bodies (39). Qualitatively, ATII cells from mock-infected mice contained large, homogeneous, rounded mitochondria with darker (more electron-dense) cristae protruding into the mitochondrial matrix (Fig.

2.1A). In comparison, ATII cells from IAV-infected mice contained mitochondria that varied greatly in shape and size but were uniformly highly electron-dense (Fig 2.1B).

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ImageJ was used to quantify characteristics of ATII cell mitochondria in TEM images.

Although the number of mitochondria in ATII cells did not change between mock- and

IAV-infected mice at 6 dpi (Fig 2.1C), mitochondria were smaller on average in ATII cells from IAV-infected mice (Fig 2.1D). Additionally, in IAV-infected mice at 6 dpi, total mitochondrial area accounted for a smaller percentage of ATII cell area (Fig. 2.1E).

These image analyses indicated quantifiable changes in the gross morphology of ATII cell mitochondria following 6 days of IAV infection and the development of ARDS.

IAV infection alters mitochondrial ROS (mtROS) and mitochondrial mass in ATII cells. Multiple mitochondrial processes generate mtROS, which can oxidize mitochondrial membrane lipids and proteins and damage the organelle (40). Using

MitoSOX, a flow cytometry dye that fluoresces red when it is oxidized by mtROS, we examined mtROS production in ATII cells at 6 dpi. Only 35% of ATII cells isolated from mock-infected mice at 6 dpi had high enough mtROS levels to oxidize MitoSOX (Fig.

2.2A). 49% of ATII cells from IAV-infected mice at 6 dpi, a significantly increased percentage, contained sufficient mtROS (Fig. 2.2A). However, the mean channel fluorescence (MCF) of MitoSox, an indicator of the magnitude of oxidation, is not significantly different between groups (Fig. 2.2B), suggesting that although more ATII cells have detectable mtROS levels following 6 days of IAV infection, the level of mtROS in each cell does not increase.

MitoTracker is another mitochondrial-permeable fluorescent dye, which accumulates in mitochondria and is used to measure mitochondrial mass. The MCF of

MitoTracker indicates the amount of dye that has accumulated in the mitochondria of a given cell. Interestingly, ATII cells isolated from IAV-infected mice at 6 dpi have a

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significantly higher MitoTracker MCF than cells isolated from mock-infected mice (Fig.

2.2C), indicating that they have greater mitochondrial mass.

IAV infection alters cardiolipin (CL) synthesis and remodeling in ATII cells. Our previous research showed that IAV infection reduced levels of phospholipids in ATII cells at 6 dpi (15). This includes phospholipids that are critical for mitochondrial membrane formation, such as PC and PE. To assess impact of IAV infection on mitochondrial membranes phospholipids specifically, we focused on CL, a phospholipid that is exclusively incorporated into mitochondria. Nonyl acridine orange (NAO) is a dye that binds to CL moieties in live cells and emits a fluorescent signal, so that CL content can be quantified by flow cytometry (37). ATII cells isolated from IAV-infected mice at 6 dpi had a significantly reduced MCF for NAO, when compared to ATII cells isolated from mock-infected mice (Fig. 2.3A). This indicates that ATII cells contain less CL following 6 days of IAV infection. To assess defects in CL synthesis, we examined gene expression of two enzymes involved in the synthesis pathway of this phospholipid. Gene expression of cardiolipin synthase (CRLS1) increased 3-fold in ATII cells at 6 days post IAV infection, compared to ATII cells at 6 days post mock infection (Fig. 2.3B). Gene expression of tafazzin (TAZ), which plays a major role in CL remodeling, was not altered by infection (Fig. 2.3B), although protein expression of this enzyme was significantly decreased (Fig. 2.3C and D).

IAV infection activates mitochondrial biogenesis in ATII cells. Changes in phospholipid availability may result in mitochondrial membrane abnormalities, if the preferred building blocks for these bilayers are present at reduced levels or unavailable.

Mitochondrial biogenesis is an important component of mitochondrial quality control,

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which may be activated to minimize or repair mitochondrial defects (41, 42). Nuclear transcription factor peroxisome proliferator-activated receptor gamma coactivator 1a

(PGC-1a) is the master regulator of mitochondrial biogenesis, which encompasses mitochondrial fusion and fission (43). Protein expression of PGC-1a was significantly increased at 6 dpi in ATII cells from infected mice (Fig. 2.4A and B). This suggests an increase in mitochondrial biogenesis, which was supported by the observed increase in mitochondrial fusion. This process is coordinated by two proteins embedded in the outer mitochondrial membrane, mitofusin 1 and 2 (MFN1 and MFN2). Gene expression of both

MFN1 and MFN2 was increased about 3-fold in live ATII cells isolated from IAV-infected mice at 6 dpi, compared to mock-infected controls. (Figure 2.4C).

IAV infection reduces mitochondrial membrane potential in ATII cells but does not induce transition pore opening. Mitochondrial membrane potential (ΔΨm) is generated as a result of the movement of electrons and protons through the ETC embedded in the

IMM. Cationic cyanine dyes such as DiIC1(5) accumulate in mitochondria with active

ΔΨm (44). In ATII cells isolated from mock-infected mice, DiIC1(5) staining revealed two populations by flow cytometry. One population, in cells with high DiIC1(5) fluorescence, had active ΔΨm. The second population, in cells with low DiIC1(5) fluorescence, had low

ΔΨm, indicating mitochondrial depolarization. Figure 2.5A shows that in mock-infected mice the majority of ATII cells had high DiIC1(5) and active ΔΨm, while only about 22% were depolarized. The percentage of ATII cells with low ΔΨm increased significantly in

IAV-infected mice. About 36% of ATII cells had depolarized mitochondria under these conditions (Fig. 2.5B and C).

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Mitochondrial depolarization indicates collapse of ETC function, and is often linked with opening of the mitochondrial permeability transition pore (mPTP) during the commitment of cells to apoptosis (45, 46). To assess this relationship in our model, we used a flow cytometric method that loads mitochondria with calcein AM dye and then chemically induces mPTP opening with ionomycin (47, 48). Calcein AM signal is limited to mitochondria by quenching the cytosolic signal with CoCl2, but the dye will leak back out to the cytosol and dissipate the mitochondrial signal if the mPTP is continuously open, as is the case in dying cells (47, 48). In murine ATII cells, calcein AM loading of mitochondria resulted in a strong fluorescent signal, which was almost completely dissipated following treatment with ionomycin to open the mPTP (Fig. 2.5D and E). In

ATII cells from mock-infected mice, ionomycin treatment decreased mean channel fluorescence (MCF) by 78% (Fig. 2.5F). Ionomycin treatment decreased MCF by 68% in

ATII cells from IAV-infected mice at 6 dpi (Fig. 2.5F). The extent of MCF reduction was not significantly different between groups, indicating that IAV infection did not induce mPTP activation in ATII cells beyond homeostatic levels.

IAV infection slows ATII cell oxygen consumption and alters mitochondrial energy production. In addition to examining ΔΨm, mitochondrial function can also be measured by determining the oxygen consumption rate (OCR) of a cell (49). Oxidative phosphorylation in the mitochondrial matrix to drive ATP production relies on the reduction of oxygen to water. OCR is therefore correlated to mitochondrial energy production. In addition, pharmacological inhibitors that target various components of the

ETC will have an impact on OCR, and therefore provide information about different parameters of mitochondrial energy production. Oligomycin inhibits ATP synthesis, and

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the difference between basal OCR and the level of OCR that occurs when ATP synthesis is halted indicates how much OCR was being used to drive this process.

Carbonilcyanide p-triflouromethoxyphenylhydrazone (FCCP) uncouples oxidative phosphorylation from the generation of a proton gradient by the ETC (50), allowing it to proceed much more rapidly and revealing the spare respiratory capacity of a cell. Finally, dual treatment with rotenone, a complex I inhibitor, and antimycin A, a complex III inhibitor (51), terminates ETC use of oxygen for oxidative phosphorylation, revealing how much OCR is providing oxygen for other cellular processes.

Figure 2.6A shows the fluctuations in OCR that occurred in ATII cells with the addition of each of these compounds. Although this graph illustrates the OCR of ATII cells from a single mock- and IAV-infected mouse at 6 dpi, the difference in OCR is immediately remarkable. Quantification of ATII cell OCR during this assay reveals that

ATII cells from IAV-infected mice had significantly lower OCRs under both basal conditions and following oligomycin treatment (Fig. 2.6B). When this OCR data was analyzed using parameters for the different components of respiration, we found that

ATII cells isolated from IAV-infected mice at 6 dpi had significantly reduced rates of ATP synthesis, although their spare respiratory capacity was not significantly diminished (Fig.

2.6C). Altogether, this data clearly indicates a reduction in mitochondrial ATP production by ATII cells during IAV infection.

IAV infection does not substantially alter electron transport chain enzyme expression in ATII cells. To begin to assess the mechanisms leading to reduced ATP production by mitochondria from infected IAV mice, we examined the gene expression of the enzyme subunits in complexes I-IV of the ETC. The majority of these genes are

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encoded in the nuclear genome (52). In live ATII cells isolated from IAV-infected mice at

6 dpi, expression of very few subunits was significantly different from expression in mock-infected mice (Fig. 3.3). This suggests that changes in mitochondrial energy production are not based on genomic alterations.

2.5 Discussion

Our results indicate that acutely lethal infection with IAV changes ATII cell mitochondrial physiology in mice. Electron microscopy revealed that mitochondria had visually altered ultrastructure at 6 dpi, with increased electron density and variations in size and shape that were not observed in ATII cells from mock-infected mice (Fig 2.1).

Electron microscopy image analysis revealed that while the number of mitochondria in

ATII cells does not change with IAV infection, the mitochondria from infected mice are significantly smaller, and take up less area within the cell. Other changes include an increase in mitochondrial mass and more ATII cells with mtROS, as well as a decrease in CL content. Finally, gene expression analysis indicates increased expression of PGC-

1a and the mitofusins, suggesting that ATII cells may upregulate mitochondrial fusion, an important quality control mechanism, during severe influenza.

The reduction in mitochondrial size and changes in mitochondrial ultrastructure support our hypothesis that altered phospholipid synthesis in ATII cells would affect mitochondria. PC and PE are critical for the formation of phospholipid bilayers for mitochondrial membranes, and the synthesis of these molecules is disrupted in ATII cells from IAV-infected mice (15). Without these essential building blocks, it would be difficult to maintain normal mitochondria. This may result in smaller mitochondria, as we

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observed at 6 dpi (Fig. 2.1). However, this conclusion may be contradicted by our flow cytometry data indicating that ATII cells have increased mitochondrial mass at 6 dpi (Fig.

2.2). MitoTracker FM Red is a rosamine dye, and its accumulation within cells is dependent on ΔΨm, which is generated by ETC activity (53). It is possible that the observed increase in mitochondrial mass following infection using this method reflects changes in ΔΨm more than it reflects their size, although we observed a reduction in

ΔΨm and ETC activity in ATII cells from infected mice (Fig. 2.6). Alternative dyes are available to study mitochondrial mass by flow cytometry [reviewed in (53)], but their use in our research is limited by the high level of background autofluorescence in ATII cells

(54).

NAO has also been used to measure mitochondria mass, as its accumulation within mitochondria is not dependent on membrane potential (53, 55, 56), but instead on

CL content. However, our data that IAV disrupts phospholipid synthesis in ATII cells cautions against the use NAO for this purpose. We showed in Fig. 2.3 that IAV infection reduces NAO fluorescence in ATII cells compared to mock-infected experimental controls. This is consistent with our previous work showing that phosphatidylglycerol

(PG) levels are reduced in ATII cells from infected mice (15). PG is the immediate precursor of nascent CL, and is converted to CL by cardiolipin synthase (CLRS) (20).

Interestingly, gene expression of CRLS is increased in live ATII cells from infected mice at 6 dpi (Fig. 2.3), suggesting that this pathway may be upregulated in an attempt to compensate for the lack of available substrate. Expression of tafazzin (TAZ), which remodels newly-synthesized CL in the mitochondria (20), does not change at the genetic level, but protein expression of this enzyme is significantly decreased at 6 dpi in ATII

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cells isolated from IAV-infected mice. This supports a defect in CL synthesis and remodeling in these cells as a result of IAV infection.

These results corroborate changes in mitochondrial size and structure that have been reported in other lung diseases. Cigarette smoke exposure induces mitochondrial fragmentation and cristae depletion in bronchial epithelial cells, and similar changes are seen in airway epithelial cells from COPD patients (22, 23). Mice exposed to chronic cigarette smoke exhibit swollen mitochondria with diminished cristae in pulmonary epithelial cells (24). ATII cells in lungs from IPF patients have increased mitochondrial content, with swollen, abnormal mitochondrial ultrastructure (25). Bronchial epithelial cells from a murine allergic asthma model also show mitochondrial swelling, along with the loss of cristae structures by electron microscopy (28). Increased mitochondrial mass and abnormal morphology have also been observed in the lungs of asthma patients (26,

27).

In addition to showing novel mitochondrial structural changes in a clinically relevant model of ARDS, we are also the first to examine the structural role of CL in this disease. We found that ATII cell mitochondria from IAV-infected mice contain significantly less cardiolipin than mitochondria from control mice (Fig 2.3). Cardiolipin is critical for inducing IMM curvature into cristae, and for the formation of respiratory supercomplexes, which assemble ETC enzymes in close proximity to increase the efficiency of ATP production and reduce ROS generation (57, 58). A loss of cardiolipin therefore has the potential to not only disturb IMM structure, but also cellular energy production. Previous work has mostly looked at CL as a pro-inflammatory DAMP and disruptor of pulmonary surfactant function once it is secreted into the extracellular space

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during acute lung injury (59, 60); we are the first to report an intercellular role for this phospholipid during acute respiratory failure.

Huang et al. observed increased expression of lysocardiolipin acyltransferase

(LYCAT), also known as acyl:CoA LYCAT (ALCAT1), a cardiolipin remodeling enzyme active in the endoplasmic reticulum and mitochondrial associated membrane (20), in lung tissue from IPF patients and in the pulmonary epithelium of mice with experimentally-induced fibrosis (61). Interestingly, over-expression of LYCAT in the lung prior to fibrotic insult reduced the severity of fibrosis in these models, suggesting a protective role for CL in alveolar epithelium in this disease (61). Although we see decreased expression of the CL remodeling enzyme TAZ in ATII cells following infection, we do see increased expression of CRLS, suggesting the upregulation of CL synthesis as a potentially protective mechanism in viral infection of the lung.

The mitochondria are a site of substantial ROS production due to the number of oxidation-reduction reactions that take place. mtROS is recognized as a second messenger for cellular signaling, and at basal levels helps to maintain homeostasis (60,

62). Substantial increases in mtROS however can damage mitochondria by oxidizing proteins, lipids and DNA (62, 63). Additionally, mtROS can act as a pro-inflammatory

DAMP, and is known to contribute to the pathogenesis of many chronic respiratory diseases [reviewed in (60, 64)]. We found that although more ATII cells from infected mice have detectable mtROS at 6 dpi, the magnitude does not increase above the mtROS levels in ATII cells from mock-infected mice (Fig. 2.2). This suggests an increase in the extent of mtROS-mediated signaling in ATII cells from infected mice, but not a pathological increase in the amount of mtROS in each cell. Elevated ROS levels have

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been observed in patients with ARDS [reviewed in (62)]. This increase may be driven by other cell types, such as neutrophils or macrophages, as mtROS is not increased above basal levels in ATII cells from mice with IAV-induced ARDS.

Another area where we observed changes was mitochondrial quality control.

Expression of PGC-1α, the master regulator of mitochondrial biogenesis was increased in ATII cells from infected mice. Expression of this transcription factor activates a cascade of genes that promote the generation of new mitochondria through fission and fusion. Consistent with our results, PGC-1α activation has been previously reported in murine models of acute lung injury (ALI) (60). Staphylococcus aureus-induced ALI increased PGC-1α expression in alveolar epithelial cells (65, 66). This work suggests that ATII cells up-regulate PGC-1α, and subsequently induce mitochondrial biogenesis to aid in resolving lung injury. In ATII cells from IAV-infected mice, gene expression of

MFN1 and MFN2 was increased three-fold (Fig. 2.3). Mitochondrial fusion facilitates the joining of mitochondria into elongated networks, which predominate in cells that rely heavily on oxidative phosphorylation for energy production, such as ATII cells (60).

Mitochondrial fusion is increased in COPD (23, 67), and MFN1 and MFN2 activity may protect against fibrosis in IPF (68). Interestingly, Chung et al. also found that deletion of

MFN1 and MFN2 in ATII cells disrupted phospholipid synthesis (68). We also observe disruptions in ATII cell phospholipid synthesis in our mouse model of ARDS, and upregulation of MFN1 and MFN2 may indicate a compensatory mechanism to address phospholipid deficiencies or reduced energy production. Mitochondrial structural abnormalities are common across chronic lung conditions, and we have shown that they also occur in an acute lung disease: ARDS. Correct mitochondrial structure is critical for

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mitochondrial function; for instance, oxidative phosphorylation relies on the ability to sequester protons in the IMS to drive ATP production. It is likely that the observed changes in mitochondrial structure disrupt mitochondrial function, which may contribute to the role of ATII cells in ARDS pathogenesis.

Our assessment of mitochondrial function in ATII cells following IAV infection found that mitochondrial energy production was significantly impaired in these cells.

Mitochondrial membrane depolarization was increased, which complemented the observed reduction in ATII cell oxygen consumption and ATP production rates. These changes did not occur as a result of ATII cell apoptosis, as IAV infection did not induce mPTP opening in ATII cells, nor did they occur due to changes in ETC complex enzyme expression.

Mitochondrial depolarization, quantified by a reduction in ΔΨm, occurs when the

ETC is unable to generate a membrane potential across the IMM. Intact ΔΨm, in comparison, is considered an indicator of healthy mitochondria. Mitochondrial depolarization has been reported in pulmonary epithelial cells in multiple lung diseases.

Chronic exposure to cigarette smoke, a model of chronic obstructive pulmonary disease

(COPD) pathogenesis significantly decreased ΔΨm in human bronchial epithelial cells, in a dose-dependent manner (69, 70). In contrast, ATII cells isolated from smoker and emphysema patient lungs had increased ΔΨm compared to ATII cells from healthy donor lungs (71). In non-small cell lung cancer, ΔΨm differs between tumor subtypes in vivo

(72). We observed an increase in the proportion of ATII cells with depolarized mitochondria in IAV-infected mice at 6 dpi (Fig. 2.5). To our knowledge, we are the first to assess ATII ΔΨm in ARDS.

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Mizumura et al. reported that mitochondrial depolarization in bronchial epithelial cells exposed to cigarette smoke extract was linked to mPTP opening, and could be reversed by cyclosporin A, which inhibits mPTP activation (69). As mitochondrial depolarization and mPTP are often linked, we used flow cytometry to assess mPTP activation in ATII cells from IAV-infected mice. mPTP opening can be quantified by determining the change in mitochondrial fluorescence in a cell after treatment with an ionophore such as ionomycin, which induces mPTP opening by triggering massive calcium efflux into the cell. Once the mPTP is opened, mitochondrial fluorescence will dissipate as the dye re-distributes throughout the cell (47, 48). If mPTP activation induces a large change in mitochondrial fluorescence, it indicates that the mPTP was not open previously. If the change in fluorescence is small, the mPTP was already active.

We found that in ATII cells from IAV-infected mice, ionomycin treatment reduced mitochondrial MCF by about 68% at 6 dpi. This was not significantly different from the reduction in mitochondrial MCF observed in ATII cells from mock-infected mice, which was about 78% (Fig. 2.5). This suggests that mitochondrial depolarization in ATII cells following IAV infection is not due to the initiation of apoptosis.

While ΔΨm provides a single measurement of mitochondrial function as it pertains to generation of a membrane potential, extracellular flux analysis using a

Seahorse analyzer permits more thorough assessment of defects in mitochondrial function. Measuring OCR as a part of extracellular flux analysis indicates how rapidly mitochondria in a given cell type are using oxygen. For example, overall oxygen consumption by healthy ATII cells, quantified by basal OCR measurement, occurs at a rate of about 100 picomoles of O2 per minute (pmol/min). In comparison, in ATII cells

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from IAV-infected mice, the basal OCR drops significantly at 6 dpi, to below 50 pmol/min. These cells are consuming oxygen much more slowly, resulting in less oxygen availability for mitochondrial functions such as oxidative phosphorylation. To determine the amount of oxygen being used for ATP production directly, we treated ATII cells with oligomycin during extracellular flux analysis. Oligomycin inhibits ATP synthase, and the resulting decrease in OCR indicates how much oxygen was being used for ATP production under basal conditions. In ATII cells from IAV-infected mice, significantly less

OCR is dedicated to ATP production at 6 dpi than in ATII cells from mock-infected mice

(Fig. 2.6).

In addition to information about real-time ATP production in cells, other parameters of OCR can also be examined using inhibitors. FCCP, a proton gradient uncoupler, allows oxidative phosphorylation to proceed at a maximal rate by un-linking this process from the generation of a proton gradient by the ETC. The subsequent increase in OCR above basal levels indicates spare respiratory capacity- the theoretical maximum rate at which ATII cells can conduct oxidative phosphorylation to produce ATP under conditions of cell stress and increased demand (73). Interestingly, there is no significant difference in spare respiratory capacity at 6 dpi between ATII cells from mock- and IAV-infected mice (Fig. 2.6).

Spare respiratory capacity is determined by a number of components, including substrate availability and ETC complex activity (73-75). The gene expression of subunits of all four ETC complexes and ATP synthase (Complex V) does not change significantly between live ATII cells isolated from mock- and IAV-infected mice at 6 dpi, with the exception of two Complex I subunit genes, NDUFS7 and NDUFS8. Additionally, protein

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expression of select ETC complex subunits does not change in ATII cells following infection (data not shown). This information, together with the substantial unused spare respiratory capacity in ATII cells, suggests that the ETC remains intact after IAV infection, and defects in oxidative phosphorylation and ATP production are driven by metabolic changes.

Reductions in mitochondrial energy production are increasingly recognized as contributing to disease pathogenesis. In the lung, reduced OCR has been reported in epithelial cells in COPD (63). ETC complex activity is reduced in ATII cells from IPF donor lungs (25), suggesting reduced ATP production in these cells. In LPS-induced models of ARDS, rabbits (76) and mice (77) were found to have lower lung ATP content and reduced mitochondrial respiration respectively. Systemically, sepsis, a major cause of ARDS (5), impairs oxidative phosphorylation and ATP production in peripheral blood mononuclear cells, and impaired mitochondrial function is associated with increased mortality risk (78, 79).

We have specifically shown reduced ATP production by oxidative phosphorylation in ATII cells in mice with IAV-induced ARDS. ATII cells are multifunctional cells with high energy demands, and defects in mitochondrial energy production may comprise their ability to maintain homeostasis in the alveoli.

Compromised ATII cell function is associated with the development of ARDS in mice

(14, 15), and mitochondrial dysfunction may underlie these changes.

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2.6 Figures

Figure 2.1: ATII cell mitochondrial ultrastructure is altered by IAV infection. A and B) Representative TEM images of ATII cells in the lungs of mock-infected (A) and IAV-infected (B) mice. Insets highlight mitochondria, denoted with white arrows. C) Number of mitochondria counted per ATII cell image captured by TEM from lungs of mock-infected and IAV-infected mice (n=3 animals per group, 10-20 cells per animal, ns: p=0.1718). D) Mean area of mitochondria measured per ATII cell image captured by TEM from lungs of mock-infected and IAV-infected mice (n=3 animals per group, 10-20 cells per animal, *: p=0.0447). E) Total mitochondrial area measured per ATII cell as a percentage of total cell area measurement of ATII cell image captured by TEM from lungs of mock-infected and IAV-infected mice (n=3 animals per group, 10-20 cells per animal, **: p<0.0001). Data are presented as mean ± SEM.

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Figure 2.2: IAV infection alters mitochondrial ROS and mitochondrial mass in ATII cells.

A) Percentage of ATII cells containing MitoSOX Red, a fluorescent dye detecting mtROS (n=6 mice per group, *: p=0.0044). B) Mean channel fluorescence (MCF) of ATII cells stained with MitoSOX Red (n=6 mice per group, ns: p=0.1855). C) MCF of ATII cells stained with MitoTracker RED FM, a fluorescent dye that measures mitochondrial mass (n=6 mice per group, *: p=0.0084). Data are presented as mean ± SEM.

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Figure 2.3: IAV infection alters cardiolipin synthesis and remodeling in ATII cells.

A) NAO MCF in ATII cells from IAV-infected mice at 6 dpi as a percentage of NAO MCF in mock-infected controls from the same experiment (n=6 mice per group, *: p=0.0109). B) Fold change in gene expression of cardiolipin synthase (CRLS) and tafazzin (TAZ) in live ATII cells from IAV-infected mice at 6 dpi, normalized to gene expression in live ATII cells from mock-infected controls (n=3 mice per group). C) Mean density ratio of tafazzin protein expression by western blot from mock- and IAV-infected mice at 6 dpi, compared to β-actin expression in mock- and IAV-infected mice at 6 dpi (n=3 mice per group, *: p=0.03). D) Representative western blot membrane evaluating tafazzin and β-actin protein expression in ATII cells isolated from mock- and IAV-infected mice at 6 dpi. Data are presented as mean ± SEM.

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Figure 2.4: IAV infection activates mitochondrial biogenesis in ATII cells.

A) Mean density ratio of PGC-1α expression by western blot of ATII cells isolated from mock- and IAV-infected mice at 6 dpi, compared to β-actin expression (n=3 mice per group, *: p=0.0317). B) Representative western blot membrane evaluating PGC-1α and β-actin protein expression in ATII cells isolated from mock- and IAV-infected mice at 6 dpi. C) Fold change in gene expression of mitofusin 1 (MFN1) and mitofusin 2 (MFN2) in live ATII cells from IAV-infected mice at 6 dpi, normalized to gene expression in live ATII cells from mock-infected controls (n=3 mice per group). Data are presented as mean ± SEM.

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Figure 2.5: IAV infection reduces mitochondrial membrane potential in ATII cells but does not induce transition pore opening.

A and B) Representative histograms for DiIC1(5) fluorescence in ATII cells isolated from mock (A) and IAV (B) infected mice at 6 dpi, denoting DiIC1(5) peaks indicating high and low mitochondrial membrane potential (ΔΨM). C) Percentage of ATII cells with low ΔΨM based on DiIC1(5) fluorescence levels (n=6-7 mice per group, *: p=0.0118). D and E) representative histograms for quantification of ATII cell mitochondrial permeability transition pore (mPTP) activation. (D) shows baseline mitochondrial fluorescence while (E) shows remaining fluorescence after activation of mPTP by ionomycin. F) Quantification of the percent change in MCF (% change MCF) from baseline after mPTP activation is chemically induced (n=5-6 per group, ns: p=0.1118). Data are presented as mean ± SEM.

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Figure 2.6: IAV infection slows ATII cell oxygen consumption and alters mitochondrial energy production.

A) Representative measurements of ATII cell oxygen consumption rate (OCR) over the timecourse of a Mito Stress Test assay for ATII cells from a mock (blue) and IAV (red) infected mouse at 6 dpi. B) Quantification of mean ATII OCR before the addition of inhibitors (BASAL) and following the addition of oligomycin (+OLIGO) and FCCP (+FCCP) (n>6 per group, #: p<0.01). C) Quantification of mean ATII OCR for different components of mitochondrial functions: basal, ATP synthesis (ATP), and spare respiratory capacity (SPARE) (n>6 per group, #: p<0.01). Data are presented as mean ± SEM.

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Figure 2.7: IAV infection does not substantially alter electron transport chain gene expression in ATII cells.

Volcano plot representing the fold changes in gene expression for components of electron transport chain complexes I-IV and ATP synthase in live ATII cells isolated from IAV-infected mice at 6 dpi. Green points represent genes down-regulated more than 2- fold. Red points represent genes up-regulated more than 2-fold. Normalized to gene expression in live ATII cells isolated from mock-infected mice, with β-actin as an endogenous control (n=3 mice/group).

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Chapter 3. Acutely Lethal Influenza Infection Alters ATII Cell Bioenergetic Metabolism

3.1 Abstract

Influenza A virus (IAV) is a respiratory pathogen that can cause critical illness and death. Up to 50% of patients hospitalized in an intensive care unit for influenza treatment may develop acute respiratory distress syndrome (ARDS), a highly lethal form of respiratory failure. In the lower airways and alveoli, IAV primarily infects alveolar type

II (ATII) cells, which play an important role in lung homeostasis. ATII cell damage is known to contribute to the development of ARDS. ATII cells from mice with IAV-induced

ARDS have altered phospholipid metabolism, and IAV infection is associated with changes in glucose metabolism. Therefore, we hypothesized that IAV infection would cause global changes in bioenergetic metabolic pathways in ATII cells. We found that

IAV infection altered glucose regulation, but not ATP production by glycolysis. In comparison, mitochondrial energy production was substantially affected, with major changes in tricarboxylic acid (TCA) cycle flux and a significant reduction in mitochondrial

ATP production. ATII cells perform a number of energetically expensive processes, and reduced ATP availability may contribute to the ATII cell dysfunction observed in IAV- induced ARDS.

3.2 Introduction

Influenza A virus (IAV) causes both seasonal influenza epidemics and global pandemics (1, 2). Although seasonal influenza is often mild, the World Health

Organization estimates that 3-5 million severe influenza cases occur each year, resulting in 250-500,000 deaths (3). Additionally, novel IAV strains have caused outbreaks with

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exceedingly high mortality rates (4, 5). Mortality from IAV often occurs secondary to the development of acute respiratory distress syndrome (ARDS), a condition characterized by the acute onset of hypoxemia, pulmonary edema, reduced lung compliance, and reduced gas exchange (6). Influenza is the leading viral cause of ARDS in humans (7), and up to 50% of influenza patients being treated in an intensive care unit (ICU) will develop ARDS (8). Once developed, the prognosis for ARDS is very poor, with in- hospital mortality around 40% and treatment options limited to supportive care such as mechanical ventilation (9, 10).

Progression of influenza to severe illness and ARDS is associated with infection of the lower respiratory tract, including the alveoli, the functional units of the lung.

Alveolar type II (ATII) epithelial cells are the target cell for IAV replication in the distal lung (11). ATII cells are also dynamic epithelial cells that maintain alveolar homeostasis to facilitate breathing. ATII cells synthesize and secrete the components of pulmonary surfactant, a film that reduces surface tension to prevent alveolar collapse during breathing (12). They also contribute to alveolar fluid clearance (AFC), maintain epithelial barrier integrity, and coordinate lung immune responses (13-17). We have previously shown that ARDS development in a mouse model of lethal IAV infection is correlated with reduced ATII cell function, including reduced AFC, reduced pulmonary surfactant protein synthesis, and altered pulmonary surfactant lipid synthesis (18-20).

Our recent work on pulmonary surfactant production included a semi-quantitative metabolomic analysis of ATII cells. We found significant changes in the availability of many surfactant phospholipids at 6 days post-inoculation (dpi) with IAV (20). Given that these phospholipids also play a major role in mitochondrial function (21), and that

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influenza infection has been reported to increase glucose uptake in cell culture models and in patient lungs (22, 23), we hypothesized that IAV infection would lead to global changes in ATII cell bioenergetic metabolism. We examined the major metabolic pathways leading to ATP production in ATII cells, glycolysis and the tricarboxylic acid

(TCA) cycle (24, 25), to identify changes following acutely lethal IAV infection.

3.3 Materials and Methods

Animal experiments. All animal experiments complied with the NRC Guide for the Care and Use of Laboratory Animals and were approved by The Ohio State University’s

Institutional Animal Care and Use Committee.

Mouse infection. 8-12 week old female C57BL/6 mice (Charles River Laboratories,

Ashland, OH, USA) were inoculated intranasally with 10,000 plaque-forming units (pfu) of influenza A/WSN/33 (H1N1) virus in 50 microliters (μl) phosphate-buffered saline

(PBS) with 0.1% bovine serum albumin (BSA) under ketamine/xylazine anesthesia (19).

This inoculum results in 100% mortality by 8 days post-inoculation (dpi), without replication in the brain (26-28). Control mice were “mock” inoculated with 50 μl PBS with

0.1% BSA under ketamine/xylazine anesthesia to mimic the route of viral infection.

Immediately preceding inoculation, mice were individually numbered and weighed. Mice were re-weighed every two days following infection to confirm disease progression.

ATII cell isolation. ATII cells were isolated from mice using a standard negative magnetic selection procedure developed to account for the unique characteristics of an influenza-infected lung. The full procedure is reviewed in Chapter 2.

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Quantitative real-time PCR (qRT-PCR). RNA was extracted from ATII cells using

Qiazol and the miRNeasy Mini Kit (both Qiagen). RNA quality and quantity were examined by spectrophotometry (NanoDrop2000). cDNA was synthesized using the RT2

PreAMP cDNA Synthesis Kit and RT2 PreAMP Pathway Primer Mixes (both Qiagen), to transcripts for genes included in the corresponding array plate. PCR arrays were run using RT2 Profiler PCR Arrays with RT2 SYBR Green qPCR Mastermix (both Qiagen).

The Mouse Glucose Metabolism PCR Array (Qiagen) was used to examine genes involved in glycolysis, including hexokinase III (HK3), phosphoenolpyruvate carboxykinase 1 (PCK1), aldolase c (ALDC), and pyruvate dehydrogenase kinase 1

(PDK1). All qRT-PCR experiments were run on a 96-well StepOnePlus Real-Time PCR

System (Applied Biosystems) using the associated StepOnePlus software. CT values were exported to Microsoft Excel and gene expression patterns were evaluated using the ΔΔCT method (29). For PCR arrays, β-actin was included on the array plate and used as a housekeeping gene.

Western blotting. For relative quantification of protein expression, freshly isolated ATII cells were vortexed then frozen at -80°C in 1:10 diluted Cell Lysis Buffer (Cell Signaling) containing 1 μL/mL benzonase nuclease (Sigma Aldrich). Once frozen overnight, cell

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lysates were thawed on ice and vortexed at maximum speed in a benchtop centrifuge at

4°C for 10 minutes. Supernatants were transferred to new 1.5 mL microcentrifuge tubes and stored on ice. To determine protein concentration, BCA assays were performed according to manufacturer instructions (Pierce BCA Protein Assay Kit) and read on a colorometric plate reader at 562 nm. Based on BCA results, protein concentrations were calculated from the generated standard curve. To prepare samples for SDS-PAGE, 10 ug of protein was mixed with appropriate volume of 4x Bolt LDS Sample Buffer and 10x

Bolt Reducing Agent (both Life Technologies), as well as PBS to achieve a uniform loading volume across samples. Samples were loaded into Bolt 4-12% Bis Tris Plus gels in the Invitrogen Mini Gel Tank with 1:20 diluted 20x Bolt MOPS SDS Running Buffer (all

Life Technologies). A dual color/fluorescent molecular weight ladder (GE Healthcare) was also loaded onto the gel, which was run at 150 volts for about 1 hour, until dye front had migrated to the bottom of the gel. Protein was transferred onto an Immobilon-P

PVDF membrane (Millipore) using the Invitrogen Mini Blot Module and diluted Bolt 20x

Transfer Buffer with 20% methanol and Bolt Antioxidant (all Life Technologies), following a standard membrane transfer protocol.

For band visualization, membranes were blocked overnight with rocking at 4°C with Blocker FL Fluorescent Blocking Buffer (Thermo Scientific). Primary antibodies against citrate synthase (1:2000), isocitrate dehydrogenase 3a (1:1000) and β-actin

(1:2000) were diluted to working concentrations in 5% BSA in TBS with 0.1% Tween

(TBS-T), and incubated overnight with rocking at 4°C. The following day, the primary antibody solution was discarded, and the membrane washed in TBS-T 3 consecutive times for 10 minutes each. Membranes were then incubated on a rocker at room

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temperature for 1 hour with the appropriate Cy5-conjugated secondary antibody (anti- mouse or anti-goat; Thermo Fisher) diluted 1:5000 in 5% BSA in TBS-T. Following 3 additional washes with TBS-T, membranes were transferred to a piece of thick filter paper and dried at 37°C for 5-10 minutes. Once dry, membranes were imaged on an

Amersham Typhoon using the Cy5 670BP30 laser, which picks up the least background fluorescence on PVDF membranes. Quantification of bands was performed on .tif image files captured by the Typhoon using ImageJ software and a standard procedure.

Expression of β-actin was used for normalization.

Analysis of the ATII cell metabolome. Metabolomic analyses were conducted at

Metabolon (Durham, NC), previously described (20). Briefly, ATII cell pellets were disrupted using a GenoGrinder (OPS Diagnostics, Lebanon, NJ) at 675 strokes/min for 2 min and then subjected to methanol extraction. Extracts were split into four aliquots and processed for analysis by UHPLC/MS in the positive, negative, or polar ion mode.

Metabolites were identified by automated comparison of ion features (retention index and accurate mass match) to a reference library of authentic chemical standards followed by visual inspection for quality control. Each ion peak was quantified by a proprietary method. Data were normalized to sample lysate Bradford protein concentration for statistical analysis.

Measuring ATII cell extracellular acidification rate (ECAR). ATII cell ECAR was measured using a Seahorse XFe24 Bioanalyzer (Agilent). ATII cells were plated in specialized 24-well Seahorse cell culture microplates (Agilent) at a density of 1×105 cells per well, in warm pH 7.4 DMEM with 1.0 M glucose, 100 mM pyruvate, 200 mM glutamine (all Agilent). Before plating, microplates were coated with 22.4 μg/mL Cell-Tak

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cell and tissue adhesive (Corning). Once cells were plated, the plate was centrifuged at

300 rpm for 2 minutes with no braking to immobilize cells in Cell-Tak coating. The microplate was then incubated without CO2 at 37°C for 30 minutes to promote cell adherence.

For 24 hours prior to Seahorse assay, a sensor cartridge was hydrated by incubating without CO2 37°C with Seahorse XF Calibrant solution (both Agilent). Once

ATII cells were adhered to the microplate, the sensor cartridge was loaded into the

Seahorse analyzer for calibration. Once calibration was complete, the microplate containing ATII cells was loaded into the analyzer for data collection using Wave software (Agilent). Each experiment consisted of a series of ECAR measurements, occurring every 3 minutes over a 1-2 hour period. Once complete, readings were exported to Microsoft Excel and the mean ECCR for each sample under each condition was determined, with standard error.

Determining ATP production rate. ATII cells were plated at a density of 8×104 cells per well in a specialized Seahorse XFp 8-well plate in warm pH 7.4 DMEM with 1.0 M glucose, 100 mM pyruvate, 200 mM glutamine (all Agilent). Before plating, microplates were coated with 22.4 μg/mL Cell-Tak cell and tissue adhesive (Corning). Once cells were plated, the plate was centrifuged at 300 rpm for 2 minutes with no braking to immobilize cells in Cell-Tak coating. The microplate was then incubated without CO2 at

37°C for 45 minutes to promote cell adherence.

For 24 hours prior to Seahorse assay, a sensor cartridge was hydrated by incubating without CO2 37°C with Seahorse XF Calibrant solution (both Agilent).

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While cells were adhering to the microplate, the sensor cartridge was removed from the incubator, and XFp ATP Rate Assay Kit (Agilent) compounds were resuspended and loaded into cartridge injector ports at the following concentrations: 1.5

µM oligomycin (glycolysis inhibitor) and 0.5 µM rotenone/antimycin A (ETC inhibitors).

The sensor cartridge was then loaded into the Seahorse XFp analyzer for calibration.

Once calibration was complete, the microplate containing ATII cells was loaded into the analyzer for data collection using Wave software (Agilent). Each experiment consisted of a series of OCR and ECAR measurements, starting with 3 basal measurements, followed by injection of oligomycin into each well, mixing, 3 measurements, then injection of rotenone/antimycin A, mixing, and 3 measurements. Once complete, readings were exported to Microsoft Excel and the Seahorse Real Time ATP Rate Assay

Report Generator (Agilent) was used to calculate oxidative phosphorylation and glycolysis contributions to ATP synthesis.

Measuring ATII cell substrate usage. Following isolation, ATII cells were plated at a density of 3×104 cells per well in a specialized Biolog plate pre-coated in cellular energy substrates (Biolog). 30 µL assay mix, containing 2X Biolog Mitochondrial Assay Solution,

6X Redox Dye MC (both Biolog), 24x saponin (30 μg/mL) and sterile water, was added to each well, and the plate was gently mixed on a plate vortex and incubated at 37°C for

1 hour. All plates (1 per sample) were then loaded into an OmniLog kinetic plate reader, which took optical density 590 (OD590) measurements every 10 minutes for 12 hours.

Once measurement period was complete, OmniLog software was used to determine the mean height of the curve for uptake of each energy substrate.

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3.4 Results

IAV infection alters ATII cell glucose metabolism, but not glycolytic flux. Aerobic glycolysis contributes substantially to cellular ATP production by producing pyruvate, which is transported into the mitochondrial matrix by the mitochondrial pyruvate carrier

(30). Once in the mitochondrial matrix, pyruvate is converted to acetyl-coA by pyruvate dehydrogenase (PDH) and enters the TCA cycle. Given the role of glycolysis in driving

ATP production in mitochondria under aerobic conditions, and reports that IAV infection increases glucose uptake in vitro and in vivo (22, 23), we investigated how IAV infection affects glycolysis in ATII cells.

We found significant changes in the gene expression of multiple enzymes that determine the fate of intracellular glucose. Gene expression of hexokinase III (HK3), the enzyme that initiates glycolysis by converting glucose to glucose-6-phosphate, increased

5-fold in live ATII cells from IAV-infected mice at 6 dpi (Figure 3.1A). Gene expression of phosphoenolpyruvate carboxykinase 1 (PCK1), the regulatory enzyme in

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gluconeogenesis, increased 63-fold (Fig. 3.1A). Gluconeogenesis is the de novo production of glucose in mammalian cells (31). In comparison, gene expression of aldolase C (ALDC) decreased 25-fold (Fig. 3.1A). In glycolysis aldolase converts 6 carbon fructose 1,6-bisphosphate into two 3 carbon sugars: glyceraldehyde-3-phosphate and dihydroxyacetone phosphate (32). It catalyzes the reverse condensation reaction during gluconeogenesis (33). Finally, gene expression of pyruvate dehydrogenase kinase 1 (PDK1) was decreased 9-fold (Fig. 3.1A). PDK1 inhibits PDH to reduce pyruvate conversion to acetyl-coA and plays a role in modulating energy homeostasis

(34). Levels of glycolytic metabolites were also altered in ATII cells from IAV-infected mice at 6 dpi. Metabolomic analysis indicated significantly reduced levels of, in the order they are generated during glycolysis: fructose-6-phosphate, fructose 1,6-bisphosphate,

3-phosphoglycerate, and phosphoenolpyruvate (Fig. 3.1B).

Interestingly, these changes in gene expression and metabolite concentration were not correlated with changes in glycolytic flux in ATII cells from IAV-infected mice at

6 dpi. The extra-cellular acidification rate (ECAR) of ATII cells measures the rate at which lactate is produced during anaerobic glycolysis. During anaerobic glycolysis, pyruvate is converted to lactate instead of entering the TCA cycle. IAV infection did not significantly increase ATII cell ECAR at 6 dpi (Fig. 3.1C). Additionally, inhibition of mitochondrial ATP production by oligomycin did not increase ATII cell ECAR in either mock- or IAV-infected mice at 6 dpi (Fig. 3.1D). This suggests that ATII cells do not rapidly activate anaerobic glycolysis to generate ATP in the absence of mitochondrial

ATP production, during either homeostasis or IAV infection. Finally, metabolomic analysis indicated that IAV infection did not significantly alter pyruvate or lactate levels in

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ATII cells at 6 dpi (Fig. 3.1E). This suggests that glycolytic flux into the TCA cycle or lactate production is not altered by IAV infection.

IAV infection alters TCA cycle flux in ATII cells. The TCA cycle, which occurs in the mitochondrial matrix, consists of a series of enzymatic reactions that produce electron carriers for the electron transport chain (ETC). Metabolomic analysis indicated that ATII cells from IAV-infected mice at 6 dpi contained significantly reduced levels of many TCA cycle intermediates, including citrate, alpha-ketoglutarate (α-KG), succinate, fumarate, and malate (Fig. 3.2A). Analysis of metabolite utilization by live ATII cells showed increased utilization of α-KG and reduced utilization of fumarate and malate in cells from

IAV-infected mice at 6 dpi (Fig. 3.2B). Altogether, these results suggest that metabolites are not cycling normally through the TCA cycle following infection. This is supported by

+ the finding that ATII cells had significantly reduced levels of FAD /FADH2 (FAD) at 6 dpi following IAV infection (Figure 3.2C). FAD is an important electron carrier in mitochondrial ATP production by the ETC.

IAV infection alters TCA cycle enzyme expression in ATII cells. We used gene and protein expression techniques to assess IAV infection-induced changes that might lead to alteration in TCA cycle function. We found changes in expression of multiple TCA cycle enzymes, at both the gene and protein levels (Fig. 3.3). Gene expression of aconitase 1 (ACO1) was reduced 3-fold in live ATII cells from IAV-infected mice, and gene expression of succinyl-coA ligase, subunit beta (SUCLG2) was reduced 2.8-fold

(Fig. 3.3A). Aconitase converts citrate to isocitrate at the beginning of the TCA cycle. A subunit of succinyl-coA synthetase, SUCGL2 is involved in the conversion of succinyl- coA to succinate. Protein expression of citrate synthase (CS), which catalyzes the initial

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reaction in the TCA cycle, converting acetyl-coA to citrate, was significantly reduced in

ATII cells from IAV-infected mice at 6 dpi (Fig. 3.3B and C). Another TCA cycle enzyme, isocitrate dehydrogenase 3 alpha (IDH3A), a subunit of IDH3, which converts isocitrate to α-KG, also had significantly reduced protein expression at 6 dpi in ATII cells from IAV- infected mice (Fig. 3.3D and E).

IAV infection alters ATII cell energy production rates. Extracellular flux analysis on a

Seahorse bioanalyzer can be used to gain high-level information about cellular ATP production rates. Overall, ATII cells from IAV infected mice produced significantly lower amounts of ATP per minute than ATII cells from mock-infected mice (Fig. 3.4A). This was mostly due to a significantly reduced rate of mitochondrial ATP production (Fig.

3.4B), as the rate of ATP production by anaerobic glycolysis in ATII cells did not change significantly between IAV-infected and mock-infected mice (Fig. 3.4C). The maintenance of a steady rate of glycolytic ATP production following infection resulted in an increased percentage of ATP coming from glycolysis in ATII cells from IAV-infected mice at 6 dpi

(Fig. 3.4D). Glycolysis contributed 18% of the ATP production rate in ATII cells from IAV- infected mice, compared to 11% in mock-infected mice. This suggests that IAV infection induces a defined, but not substantial, increase in glycolysis in ATII cells.

3.5 Discussion

Our investigation of murine ATII cell bioenergetic metabolism during IAV-induced

ARDS found that energy production was impacted in several areas. The gene expression of a number of enzymes involved in glucose metabolism was significantly altered. Glycolysis, the breakdown of glucose to produce pyruvate, ATP and NADH,

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begins with the conversion of glucose to glucose-6-phosphate by hexokinase.

Mammalian cells express multiple hexokinase isoforms, and overexpression of hexokinase II (HK2) was associated with reduced cell death and preserved mitochondrial integrity during hypoxia in A549 cells, an ATII-like cell line (35). We found increased expression of a different isoform, hexokinase III (HK3) in live ATII cells from

IAV-infected mice at 6 dpi (Fig. 3.1). Unlike HK2, which localizes to mitochondria, HK3 localizes to the nuclear membrane (35), but HK3 overexpression in immortalized hepatocytes has also been associated with preservation of mitochondrial integrity, including restored mitochondrial membrane potential (ΔΨm) and increased ATP production, under hypoxic conditions (36). It is possible that increased HK3 expression in our model is also to promote mitochondrial integrity in a low-oxygen environment such as the virus-damaged alveolus experiences.

The most profound change in ATII cell gene expression that we observed was the many-fold increase in expression of phosphoenolpyruvate carboxykinase 1 (PCK1)

(Fig. 3.1). PCK1 is the key cytosolic regulatory enzyme in gluconeogenesis- the synthesis of glucose from cellular carbon substrates. In mammals, the liver is responsible for most gluconeogenesis, which does not occur in the lungs under homeostatic conditions (31, 37). This may explain the magnitude of the change in PCK1 expression in ATII cells following infection. Gluconeogenesis has been reported to occur in lung cells during lung cancer, although this was linked to increased expression of mitochondrial phosphoenolpyruvate carboxykinase 2 (PCK2) (38). Increased PCK1 expression has been observed in other tumor types, and it has been proposed the PCK1 acts as a regulator of glycolysis, gluconeogenesis and the TCA cycle (39). PCK1 and

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PCK2 regulate cataplerosis- the removal of metabolites from the TCA cycle for utilization in other cellular processes, and overexpression of PCK1 in glucose-starved liver cancer cells induced cataplerosis, bioenergetic crisis and cell death (40).

We also observed decreased expression of some glycolysis enzymes (Fig. 3.1).

Expression of aldolase C (ALDC), was significantly reduced. ALDC is a highly conserved enzyme that converts fructose 1,6-bisphosphate to glyceraldehyde-3-phosphate and dihydroxyacetone phosphate during glycolysis and performs the reverse reaction during gluconeogenesis (32, 33). Reduced levels of ALDC could slow both of these processes.

Expression of pyruvate dehydrogenase kinase 1 (PDK1) was also reduced. PDK inactivates pyruvate dehydrogenase (PDH), which converts pyruvate to acetyl-coA for entry into the TCA cycle, connecting glycolysis to aerobic respiration. PDH inhibition by

PDK decreases glucose utilization and promotes fatty acid oxidation for energy production (34). Hypoxia has been found to increase PDK1 expression in MLE-15 cells, an ATII-like cell line (24). Additionally, PDK4, another PDK isozyme, was reported to be upregulated in mouse lungs following IAV infection, and chemical inhibition of PDK4 following infection improved survival and decreased disease severity (41, 42). It is possible that the observed down-regulation of PDK1 in our model is a mechanism to maintain PDH function and facilitate pyruvate flux into the TCA cycle. Our metabolomic data indicate that ATII cells maintain homeostatic levels of pyruvate and lactate following

IAV infection (Fig. 3.1), which suggests pyruvate metabolism is unchanged by infection.

Pyruvate produced by glycolysis has two energetic fates: it can either enter the mitochondria for conversion to acetyl-coA and participation in aerobic energy production via the ETC, or it can be converted to lactate by lactate dehydrogenase during anaerobic

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glycolysis. The extracellular acidification rate (ECAR) of media during a Seahorse assay serves to measure the rate at which this process is occurring (43). We found that the

ECAR of ATII cells isolated from IAV-infected mice at 6 dpi does not vary significantly from mock-infected mice (Fig. 4.1C). Moreover, the inhibition of aerobic ATP production by oligomycin does not induce a shift to anaerobic glycolysis in ATII cells from either mock- or IAV-infected mice (Fig. 4.1D), suggesting that ATII cells do not rely on this energy pathway for ATP production. It has long been recognized that ATII cells are highly reliant on mitochondria for ATP production to meet their functional needs (25, 44,

45), but recent work has suggested that IAV infection increases glycolysis in pulmonary epithelial cells in a manner similar to the Warburg effect (22, 23). We do not see this phenomenon in our model, which may more accurately represent the behavior of ATII cells in an infected lung, since we isolate our cells from mice following 6 days of in vivo infection and run our Seahorse assays immediately. Additionally, IAV replication peaks at 4 dpi in our model (19, 28), so we may be assessing ATII cell metabolism during a different phase on infection than most in vitro models.

The maintenance of consistent glycolytic flux following infection initially appears to be inconsistent with the significant changes in glycolytic intermediate levels observed in ATII cells following infection (Fig. 3.1). Metabolomic analysis indicates that a number of glycolysis intermediates are present at significantly reduced levels in ATII cells from

IAV-infected mice. However, when combined with the changes in the genetic regulation of glucose metabolism (Fig. 3.1), these results suggest that ATII cells initiate compensatory mechanisms to maintain glycolytic flux following IAV infection.

Additionally, it has recently been proposed that ATII cells derive a significant portion of

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their energy from fatty acid oxidation (FAO), and that induction of acute lung injury via

LPS instillation impairs FAO in ATII cells (46). In light of this, it is possible that the lack of changes in ATII cell glycolysis in response to infection reflect the minor role that this pathway plays in ATII cell energy production.

If this is the case, the TCA cycle may be more pertinent for examining ATII cell energy balance as both glycolysis and FAO feed into these reactions. Indeed, we do see significant alterations to TCA cycle metabolites in ATII cells from IAV infected mice.

Biolog assays with live ATII cells found that cells from IAV-infected mice had reduced utilization of most, but not all, TCA cycle metabolites from assay media (Fig. 3.2).

Metabolomic analysis also found altered levels of TCA cycle metabolites from IAV- infected mice (Fig. 3.3). Both methods identified a reduction in fumarate and malate, while α-KG utilization was reduced, but metabolite levels were increased in IAV-infected mice at 6 dpi. Similarly, succinate utilization was reduced while metabolite levels did not change following infection. Overall, this data suggests that TCA cycle flux is altered in

IAV-infected mice. This is supported by a reduction in ATII cell levels of FAD, an electron carrier involved in TCA cycle reactions that is also used by the ETC, following infection

(Fig. 4.2C). The observed increase in ATII cell α-KG concentration following infection may be due to anaplerosis, the synthesis of TCA cycle intermediate metabolites to promote flux through the cycle. Glutaminolysis converts glutamine to glutamate to α-KG to supply the TCA cycle during anaplerosis. This often occurs when mitochondrial citrate levels are decreased (47). This may occur in ATII cells following IAV infection as citrate synthase enzyme expression is significantly reduced in infected mice at 6 dpi (Fig. 3.3).

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A number of other TCA cycle enzymes are also differentially expressed following infection (Fig. 3.3). Gene expression of aconitase 1 (ACO1) and succinyl co-A ligase, β subunit (SUCLG2) was significantly decreased in live ATII cells from IAV-infected mice.

Aconitase reversibly converts citrate to isocitrate, and when localized to the cytosol, as is the case with ACO1, also plays a role in redox signaling and iron homeostasis (48, 49).

SUCGL2 is a subunit of succinyl-CoA synthetase, which converts succinyl-CoA to succinate (50). Furthermore, isocitrate dehydrogenase 3 alpha (IDH3A), which converts isocitrate to α-KG in the rate-limiting step of the TCA cycle, has reduced enzyme expression in ATII cells from IAV-infected mice at 6 dpi. IDH has multiple functions secondary to the TCA cycle, including a role in epigenetic regulation of gene expression, and is increasingly recognized as a driver of disease states (51). Overall, substantial changes in TCA cycle enzyme expression and metabolite availability will alter flux through the cycle, with downstream effects on mitochondrial ATP production, which is dependent on TCA cycle products.

Determination of ATP production rate by Seahorse assay provides an overarching view of ATII cell bioenergetic metabolism following IAV infection. The contributions of mitochondria and glycolysis to ATP synthesis are determined by measuring oxygen consumption rate (OCR) and ECAR before and after oligomycin inhibition of ATP synthase, and stoichiometric calculations are used to determine rates of ATP production based on collected assay data and media composition (52). Overall,

ATII cell ATP production rate is significantly reduced at 6 days post IAV-infection (Fig.

3.4). The majority of this reduction comes from a significant drop in mitochondrial ATP production, as ATP production by glycolysis does not change between uninfected and

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infected ATII cells (Fig. 3.4). As a result, ATII cells derive more ATP from glycolysis following infection, although the majority of their ATP continues to come from mitochondrial oxidative phosphorylation (Fig. 3.4). As discussed earlier, ATII cells are known to be highly reliant on mitochondria for energy production, and have three times as many mitochondria as other lung cells (44). Our data suggest that ATII cells lack the capacity to substantially increase glycolytic ATP production when mitochondrial oxidative phosphorylation is impaired. This may result in a bioenergetic crisis that leads to reduced ATII cell function, a phenotype which is associated with the development of

ARDS following IAV infection.

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3.6 Figures

Figure 3.1: IAV infection alters ATII cell glucose metabolism but not glycolytic flux.

A) Fold change in gene expression for hexokinase III (HK3), phosphoenolpyruvate carboxykinase 1 (PCK1), aldolase C (ALDC), and pyruvate dehydrogenase kinase 1 (PDK1) in live ATII cells isolated from IAV-infected mice at 6 dpi, compared to expression in live ATII cells isolated from mock-infected mice at 6 dpi. Normalized to β- actin as endogenous control (n= 3 mice per group). B) Metabolite levels of frustose-6- phosphate (F-6-P), fructose-1,6-bisphosphate (F-1,6-P), 3-phosphoglycerate (3-PG) and phosphoenolpyruvate (PEP) in ATII cells isolated from IAV- and mock-infected mice at 6 dpi (n=5-6 per group, #: p<0.01, **: p<0.001). C) Basal extracellular acidification rate (milli-pH units per minute) of media containing ATII cells isolated from mock- and IAV- infected mice at 6 dpi (n=13-20 per group, ns= not significant) D) Extracellular acidification rates (mpH/min) of media containing ATII cells isolated from mock- and IAV- infected mice at 6 dpi under basal conditions (Basal), and following injection with oligomycin (Oligo) (n=4 per group, ns= not significant). E) Metabolite levels of pyruvate and lactate in ATII cells isolated from mock- and IAV-infected mice at 6 dpi (n=5-6 per group). Data are presented as mean ± SEM.

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Figure 3.2: IAV infection alters TCA cycle flux in ATII cells

A) Utilization of TCA cycle metabolites by live ATII cells isolated from mock- and IAV- infected mice at 6 dpi over a 12 hour period (α-KG= alpha-ketoglutarate, n=4 per group, *: p<0.05, **: p< 0.001). B) Levels of TCA cycle metabolites α-ketoglutarate (α-KG), succinate (SUCC), fumarate (FUM) and malate (MAL) in ATII cells isolated from mock- + and IAV-infected mice at 6 dpi (n=5-6 per group, #: p<0.01). C) Levels of FAD /FADH2 (FAD) in ATII cells isolated from mock- and IAV-infected mice at 6 dpi (n=5-6 per group, #: p<0.01). Data are presented as mean ± SEM.

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Figure 3.3: IAV infection alters TCA cycle enzyme expression in ATII cells.

A) Fold change in gene expression of aconitase (ACO1) and succinyl-CoA ligase, subunit beta (SUCGL2) in live ATII cells from IAV-infected mice at 6 dpi. Fold change compared to expression in live ATII cells from mock-infected mice at 6 dpi and normalized to expression of β-actin as endogenous control (n= 3 per group). B) Mean density ratio of citrate synthase expression, compared to β-actin expression, by western blot of ATII cells isolated from IAV- or mock-infected mice at 6 dpi (n=3 per group, *: p=0.006). C) Representative western blot membrane evaluating citrate synthase (CS) and β-actin protein expression in ATII cells isolated from mock- and IAV-infected mice at 6 dpi. D) Mean density ratio of isocitrate dehydrogenase 3a (IDH3A) expression, compared to β-actin expression, by western blot of ATII cells isolated from IAV- or mock- infected mice at 6 dpi (n=3 per group, *: p=0.001) E) Representative western blot membrane evaluating IDH3A and β-actin protein expression in ATII cells isolated from mock- and IAV-infected mice at 6 dpi. Data are presented as mean ± SEM.

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Figure 3.4: IAV infection alters ATII cell ATP production rates.

A) Total picomoles of ATP generated per minute (pmol ATP/min) by ATII cells isolated from mock (white) and IAV (grey) infected mice at 6 dpi (n=3-4 per group, *: p=0.0346). B) Picomoles of ATP generated per minute (pmol ATP/min) by mitochondria in ATII cells isolated from mock (white) and IAV (grey) infected mice at 6 dpi (n=3-4 per group, *: p=0.0324). C) Picomoles of ATP generated per minute (pmol ATP/min) by glycolysis in ATII cells isolated from mock (white) and IAV (grey) infected mice at 6 dpi (n=3-4 per group). D and E) Portions of ATP generation contributed by mitochondria (purple) and glycolysis (pink) in live ATII cells isolated from mock and IAV infected mice at 6 dpi (n=3- 4 per group). Data are presented as mean ± SEM.

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Chapter 4. CDP-choline Rescue of Phospholipid Synthesis Improves Mitochondrial Function in Mice with IAV-induced ARDS

4.1 Abstract

Acute respiratory distress syndrome (ARDS) is a type of respiratory failure characterized by acute hypoxemia, pulmonary edema, reduced lung compliance and decreased gas exchange. Highly lethal, it is often the cause of death in cases of severe influenza, and the all-cause mortality rate hovers around 40%. Currently, ARDS can only be treated with supportive care and often requires mechanical ventilation. Most proposed pharmacological interventions for ARDS have failed in clinical trials. There is a clear need for novel therapeutic strategies that can prevent or treat ARDS. We have previously shown that development of ARDS is associated with alveolar type II (ATII) cell dysfunction in mice with severe influenza. In healthy lungs, ATII cells are responsible for maintaining alveolar homeostasis. In our model, influenza A virus (IAV) infection significantly decreases phosphatidylcholine (PC) synthesis in ATII cells. PC is a major cellular phospholipid and an important component of mitochondrial membranes. We found that restoring ATII cell PC synthesis by treating mice with the PC precursor CDP- choline following infection results in significantly improved ATII cell mitochondrial function, as well as attenuation of clinical signs of ARDS in mice. Therefore, we propose

CDP-choline as a potential therapeutic agent for the treatment of viral ARDS.

4.2 Introduction

Acute respiratory distress syndrome (ARDS) is a type of acute respiratory failure characterized by hypoxemia, pulmonary edema, reduced lung compliance and decreased gas exchange. This condition is associated with substantial morbidity and

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mortality, with approximately 40% of patients dying while hospitalized (1, 2). Survivors often struggle with respiratory insufficiency, muscle weakness and psychological sequelae that may persist for years (3). ARDS often develops secondary to lung infections, and influenza is the leading viral cause of ARDS (4).

Influenza A viruses (IAV) cause seasonal influenza epidemics and possess demonstrated pandemic potential (5-7). The World Health Organization estimates that 3-

5 million people suffer from severe influenza each year, resulting in 250-500,000 deaths

(8). As many as 50% of severe influenza cases in intensive care units (ICUs) will progress to ARDS (9). While influenza can be prevented with yearly vaccination and treated early with neuraminidase inhibitors (10-12), ARDS can only be treated with supportive care. This often requires mechanical ventilation in an ICU setting, a time- and resource-intensive process (13, 14). There is a clear and urgent need to develop novel therapeutic strategies to treat ARDS, especially as the novel coronavirus disease 2019

(COVID-19) pandemic may dramatically increase incidence of this condition (15).

We have previously shown that development of ARDS is associated with changes in alveolar type II (ATII) cell function in a mouse model of acutely lethal IAV infection (16-18). ATII cells are critical for maintaining homeostasis within the lung. They conduct active ion transport to drive alveolar fluid clearance, synthesize and secrete pulmonary surfactant to reduce surface tension and prevent alveolar collapse, and coordinate innate immune responses to pathogens (19-21). Among other changes, ATII cell production of pulmonary surfactant phospholipids is significantly altered following

IAV-infection (17). Levels of phosphatidylcholine (PC) and phosphatidylethanolamine

(PE), the major surfactant phospholipids, are significantly reduced in ATII cells at 6 days

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post-IAV infection, as are levels of their immediate precursors for de novo synthesis by the Kennedy pathway, cytidine diphosphate-choline (CDP-CHO) and CDP-ethanolamine respectively. Interestingly, PC and PE are also the major phospholipids in mitochondrial membranes (22).

Mitochondria are composed of two phospholipid bilayer membranes, the outer mitochondrial membrane (OMM) and the inner mitochondrial membrane (IMM) The

OMM contains 54% PC and 29% PE, while the IMM contains 40% PC and 34% PE, as well as 18% cardiolipin, a phospholipid specific to mitochondrial membranes (22-24).

The mitochondrial matrix, located within the IMM, is the site of many reactions that drive cellular energy production, including beta-oxidation of fatty acids, the tricarboxylic acid

(TCA) cycle and the electron transport chain (ETC) (25-27). The ETC pumps protons across the IMM to generate proton-motive force to drive ATP production by ATP synthase during oxidative phosphorylation (28, 29). Proper mitochondrial structure is therefore critical for maintaining mitochondrial function. We hypothesized that changes in

ATII cell phospholipid availability would affect mitochondria, and that restoring Kennedy pathway phospholipid synthesis would rescue mitochondrial function. We found that treatment of IAV-infected mice with CDP-CHO, the PC precursor, did indeed restore ATII mitochondrial energy production, as well as attenuating clinical signs of ARDS. This suggests that mitochondrial energy availability is critical to maintaining ATII cell function and preventing progression to respiratory failure.

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4.3 Materials and Methods

Animal experiments. All animal experiments complied with the NRC Guide for the Care and Use of Laboratory Animals and were approved by The Ohio State University’s

Institutional Animal Care and Use Committee.

Mouse infection. 8-12 week old female C57BL/6 mice (Charles River Laboratories,

Ashland, OH, USA) were inoculated intranasally with 10,000 plaque-forming units (pfu) of influenza A/WSN/33 (H1N1) virus in 50 microliters (μl) phosphate-buffered saline

(PBS) with 0.1% bovine serum albumin (BSA) under ketamine/xylazine anesthesia (18).

This inoculum results in 100% mortality by 8 days post-inoculation (dpi), without replication in the brain (30-32). Control mice were “mock” inoculated with 50 μl PBS with

0.1% BSA under ketamine/xylazine anesthesia to mimic the route of viral infection.

Immediately preceding inoculation, mice were individually numbered and weighed. Mice were re-weighed every two days following infection to confirm disease progression.

CDP-choline treatment regimen. Mice were injected interperitoneally (ip) with 100 µg

CDP-choline (CDP-CHO) in 50 µL sterile saline each day following infection (1-5 dpi) and euthanized for endpoint experiments at 6 dpi. Combination-treated mice were injected ip with 100 µg CDP-CHO in 50 µL sterile saline each day as well as 10 µg CDP- diacylglycerol (CDP-DAG) and euthanized for endpoint experiments at 6 dpi.

Transmission electron microscopy and image analysis. Whole lungs were perfusion- fixed with glutaraldehyde and prepared for transmission electron microscopy analysis by standard methods (33). Ultrastructure was visualized using a JEM-1400 transmission electron microscope (JEOL, Peabody, MA) linked to an Olympus SIS Veleta 2K camera

(Olympus Soft Imaging Solutions).

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PVDF membrane (Millipore) using the Invitrogen Mini Blot Module and diluted Bolt 20x

Transfer Buffer with 20% methanol and Bolt Antioxidant (all Life Technologies), following a standard membrane transfer protocol.

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For band visualization, membranes were blocked overnight with rocking at 4°C with Blocker FL Fluorescent Blocking Buffer (Thermo Scientific). Primary antibodies against tafazzin (1:1000) and citrate synthase (1:2000) were diluted to working concentrations in 5% BSA in TBS with 0.1% Tween (TBS-T), and incubated overnight with rocking at 4°C. The following day, the primary antibody solution was discarded and the membrane washed in TBS-T three consecutive times for 10 minutes each.

Membranes were then incubated on a rocker at room temperature for 1 hour with the appropriate Cy5-conjugated secondary antibody (anti-mouse or anti-goat; Thermo

Fisher) diluted 1:5000 in 5% BSA in TBS-T. Following three additional washes with TBS-

T, membranes were transferred to a piece of thick filter paper and dried at 37°C for 5-10 minutes. Once dry, membranes were imaged on an Amersham Typhoon using the Cy5

670BP30 laser, which picks up the least background fluorescence on PVDF membranes. Quantification of bands was performed on .tif image files captured by the

Typhoon using ImageJ software and a standard procedure. Expression of β-actin was used for normalization.

Analysis of the ATII cell metabolome. Metabolomic analyses were conducted at

Metabolon (Durham, NC), as previously described (17). Briefly, ATII cell pellets were disrupted using a GenoGrinder (OPS Diagnostics, Lebanon, NJ) at 675 strokes/min for 2 min and then subjected to methanol extraction. Extracts were split into four aliquots and processed for analysis by UHPLC/MS in the positive, negative, or polar ion mode.

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proprietary method. Data were normalized to sample lysate Bradford protein concentration for statistical analysis.

Mitochondrial membrane potential assay. ATII cell mitochondrial membrane potential was quantified on an Attune Nxt Flow Cytometer (Invitrogen) using the MitoProbe

DiIC1(5) kit (excitation/emission 633/670 nm, Invitrogen). ATII cells were incubated with

10 μM DiIC1(5) in for 30 minutes at 37°C. Positive control samples were incubated with

100 μM CCCP, a membrane potential dissipator, for 5 minutes at 37°C, prior to addition of DiIC1(5). Following incubation, the dye was removed by centrifugation at 1000 rpm for

5 minutes at 4°C. Cells were re-suspended in 1 mL PBS and read immediately on the flow cytometer using the following parameters: forward scatter (FSC) 160V, side scatter

(SSC) 340V, red laser 1 (RL1) 290V.

Measuring mitochondrial oxygen consumption rate (OCR). ATII cell OCR was measured using a Seahorse XFe24 Bioanalyzer and XFe24 Mito Stress Test Kit (both

Agilent). ATII cells were plated in specialized 24-well Seahorse cell culture microplates

(Agilent) at a density of 1×105 cells per well, in warm pH 7.4 DMEM with 1.0 M glucose,

100 mM pyruvate, 200 mM glutamine (all Agilent). Before plating, microplates were coated with 22.4 μg/mL Cell-Tak cell and tissue adhesive (Corning). Once cells were plated, the plate was centrifuged at 300 rpm for 2 minutes with no braking to immobilize cells in Cell-Tak coating. The microplate was then incubated without CO2 at 37°C for 30 minutes to promote cell adherence.

For 24 hours prior to Seahorse assay, a sensor cartridge was hydrated by incubating without CO2 37°C with Seahorse XF Calibrant solution (both Agilent). While cells were adhering to the microplate, the sensor cartridge was removed from the

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incubator, and Mito Stress Test compounds were resuspended and loaded into cartridge injector ports at the following concentrations: 1 µM oligomycin (glycolysis inhibitor), 4 µM

FCCP (proton gradient decoupler), and 0.5 µM rotenone/antimycin A (ETC inhibitors).

The sensor cartridge was then loaded into the Seahorse analyzer for calibration. Once calibration was complete, the microplate containing ATII cells was loaded into the analyzer for data collection using Wave software (Agilent). Each experiment consisted of a series of OCR measurements, starting with 3 basal measurements, followed by injection of oligomycin into each well, mixing, 3 measurements, injection of FCCP, mixing, 3 measurements, injection of rotenone/antimycin A, 3 measurements. Once complete, readings were exported to Microsoft Excel and the mean OCR for each sample under each condition was determined, with standard error.

Assessment of ARDS criteria in mice. Carotid oxygen saturation, alveolar fluid clearance, bronchoalveolar lavage, differential cell counts, and histology experiments were performed as previously described (30, 32, 34-36)

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4.4 Results

Daily CDP-choline treatment following IAV infection rescues ATII cell phosphatidyl-choline (PC) synthesis. Most cellular PC is synthesized de novo by the

Kennedy pathway (22, 37). This process begins with the uptake of dietary choline, which is converted to choline-phosphate, then combined with cytidine triphosphate (CTP) to generate CDP-choline (CDP-CHO). CDP-CHO is then combined with a diacylglycerol

(DAG) molecule to produce PC. CDP-CHO is virtually undetectable in ATII cells from

IAV-infected mice at 6 dpi, and PC levels are significantly reduced (17).

To restore movement of metabolites through the Kennedy pathway of de novo

PC synthesis, mice were treated daily with 5 mg/kg CDP-CHO ip following IAV-infection

(Figure 4.1A). This treatment regimen significantly increased the amount of PC in bronchoalveolar lavage fluid (BALF) from IAV-infected mice at 6 dpi (Fig. 4.1B), suggesting that PC synthesis was corrected. Treatment of infected mice with daily CDP-

CHO, in conjunction with daily CDP-DAG, significantly increased total ATII cell PC levels

(Fig. 4.1D), as well as levels of upstream precursors choline and choline-phosphate (Fig.

4.1C). These results suggest that exogenous CDP-choline treatment may restore PC synthesis via the Kennedy pathway.

Daily CDP-choline treatment following IAV infection improves ATII cell mitochondrial physiology. Electron microscopy images (EM) of ATII cells from IAV- infected, CDP-CHO-treated mice presented mitochondria that were less electron-dense than those in IAV-infected, untreated ATII cells (Fig. 4.2B and C). ATII cell mitochondria from treated mice also had an organized internal structure and overall morphology that more closely resembled those in ATII cells from uninfected mice (Fig 4.2A and C). ATII

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cells from CDP-CHO treated mice also had significantly higher levels of tafazzin expression by Western blot than cells from untreated mice (Fig. 4.2D and E). This suggests that cardiolipin remodeling may be restored following CDP-CHO treatment.

Daily CDP-CHO treatment improves ATII cell mitochondrial function. CDP-CHO treatment restored mitochondrial membrane potential (ΔΨm) to uninfected levels in ATII cells from IAV-infected, treated mice (Fig. 4.3A). In line with this, Mito Stress Tests of

ATII cells from CDP-CHO-treated were very similar to mock-infected mice (Fig. 4.3B), and CDP-CHO treatment significantly improved ATII cell basal OCR and ATP synthesis rates beyond mock-infected, homeostatic levels (Fig. 4.3C). Additionally, ATII cells from

IAV-infected, CDP-CHO-treated mice had normalized enzyme expression of citrate synthase at 6 dpi (Fig. 4.3D and E), suggesting improved TCA cycle function following

CDP-CHO treatment.

CDP-CHO treatment following IAV infection attenuates ARDS at 6 dpi. Mice treated with CDP-CHO for 5 days following IAV infection had significantly improved oxygen saturation (Fig. 4.4A), and normal alveolar fluid clearance rates (Fig. 4.4B). BALF from treated mice at 6 dpi showed reduced infiltration of inflammatory cells, with significantly fewer alveolar macrophages and neutrophils (Fig. 4.4C). Finally, histology from IAV- infected mice, both untreated and IAV- CDP-CHO-treated, also indicated less infiltrates and more intact alveolar architecture in the CDP-CHO-treated group following infection

(Fig. 4.4E).

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4.5 Discussion

The Kennedy pathway is a particularly useful tool for examining ATII cell phospholipid synthesis. The majority of cellular PC is produced by this mechanism, which is known to be disrupted in murine ATII cells following IAV infection (17). Our initial experiments determined that treatment with exogenous CDP-CHO following IAV infection does indeed restore PC synthesis in ATII cells (Fig. 4.1). Our treatment regimen led to a significant increase in the amount of PC in BALF obtained from IAV- infected, CDP-CHO treated mice. As BALF contains pulmonary surfactant components, this suggests increased PC synthesis and secretion as part of pulmonary surfactant, a process unique to ATII cells (21, 38). Additionally, treatment of infected mice with CDP-

CHO and CDP-DAG resulted in significantly increased levels of choline, choline- phosphate (both PC precursors upstream of CDP-CHO), and PC in ATII cells. Although we are unable to differentiate effects due to CDP-CHO from effects due to CDP-DAG in this combination regimen, these results suggest that CDP-CHO at least partially improves Kennedy pathway PC synthesis. It is interesting that restoring PC synthesis in

ATII cells from infected mice increases levels of choline and choline-phosphate, as IAV infection does not alter levels either of these upstream metabolites (Fig. 4.1). PC synthesis is thought to be regulated predominantly by the cellular localization and conformation of the enzyme the produces CDP-CHO, CTP:phosphocholine cytidylyltransferase (CCT) (39). However, it is possible that increases in cellular CDP-

CHO and/or PC following treatment initiate a positive feedback mechanism that increases choline uptake and Kennedy pathway activity.

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Treatment with CDP-CHO alone following infection induces profound changes in

ATII cell mitochondrial structure and function. ΔΨm, which is generated by ETC activity across the IMM, is restored to uninfected levels. Moreover, multiple parameters of mitochondrial respiration are significantly improved, with basal oxygen consumption and

ATP production rates even higher than those of ATII cells from mock-infected mice (Fig.

4.3). This dramatic improvement is accompanied by other changes in the mitochondrial matrix, including an increase in citrate synthase enzyme expression (Fig. 4.3). Citrate synthase is responsible for the conversion of oxaloacetate and acetyl-coA to citrate at the beginning of the TCA cycle (40), and increased enzyme expression may promote

TCA cycle activity, which would in turn generate FADH2 and NADH for the ETC.

Changes in mitochondrial function following CDP-CHO treatment are accompanied by changes in mitochondrial structure. Although not quantitative, EM images of ATII cells from IAV-infected, CDP-CHO-treated mice exhibit mitochondria that appear to have an intermediate phenotype between that of mitochondria from IAV- infected and mock-infected mice (Fig. 4.2). Mitochondrial from treated mice are less electron dense than those from IAV-infected, untreated mice and have more organized cristae that resemble mitochondria from mock-infected mice. These changes may be due to increased ATII cell PC levels following treatment. PC is a major component of mitochondrial membranes (22) and rescuing ATII cell PC levels may allow for correct mitochondrial membrane composition.

Increasing ATII cell PC synthesis with CDP-CHO treatment may also impact the production of other phospholipids by shifting cellular metabolism. Cardiolipin is a mitochondria-specific phospholipid that plays an important role in cristae formation and

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ETC function (41, 42). Cardiolipin is synthesized by cardiolipin synthase (CRLS), but undergoes substantial acyl chain remodeling by tafazzin before incorporation into mitochondrial membranes (23). CDP-CHO treatment increased enzyme expression of tafazzin in ATII cells from infected mice (Fig. 4.2). This may help restore mitochondrial structure by increasing cardiolipin remodeling and membrane incorporation, which in turn promotes cristae formation and ETC energy production.

The observed changes in mitochondrial function suggest a mechanism for how

CDP-CHO attenuates ARDS in IAV-infected mice. Mice treated with CDP-CHO following

IAV infection have significantly improved carotid arterial oxygen saturation and alveolar fluid clearance. Additionally, these mice have significantly reduced recruitment of macrophages and neutrophils to the lung, and improved lung histology (Fig. 4.4). These results are in line with previous research showing that CDP-CHO treatment improved hyperoxic lung injury in neonatal rats and demonstrating the anti-inflammatory and antioxidant characteristics of this molecule (43). The ARDS lung is a highly inflamed, reactive milieu characterized by extensive reactive oxygen species (ROS) generation, increased permeability, and pro-inflammatory molecules (44-46). In this context, it is logical that an anti-inflammatory, anti-oxidation compound could attenuate disease, although anti-inflammatory and antioxidant drugs previously proposed for ARDS treatment, including statins, glucocorticoids, and n-acetylcysteine, have all failed to improve patient outcomes (13, 47).

Our research suggests for the first time that CDP-CHO treatment also attenuates clinical signs of ARDS through an alternative mechanism. We have shown that CDP-

CHO treatment rescues mitochondrial structure and energy production in ATII cells in

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mice with acutely lethal IAV infection. Mitochondria themselves are highly susceptible to inflammation and oxidation, which induce structural and genomic damage and disrupt mitochondrial function, contributing to the pathogenesis of many lung diseases (48-50).

Reduction of mitochondrial ROS by the mitochondrial antioxidant MitoQ was recently shown to attenuate sepsis-induced acute lung injury in rats (51). MitoQ and similar compounds have also been successful in restoring mitochondrial function in laboratory models of emphysema, chronic obstructive pulmonary disease (COPD), and allergic asthma (50).

While CDP-CHO may in part improve mitochondrial function by attenuating mitochondrial damage from inflammation and oxidation, we have shown that treatment with this compound rescues ATII cell PC synthesis and improves mitochondrial structure following IAV infection (Fig. 4.1). We therefore propose that CDP-CHO rescues ATII cell mitochondrial function by providing phospholipids to correct mitochondrial membrane structure, which increases ETC function and mitochondrial ATP production. The resulting increase in cell energy availability powers ATII cell functions such as vectorial ion transport for alveolar fluid clearance, which in turn attenuates clinical signs of ARDS in mice following lethal IAV infection. This mechanism, in combination with the demonstrated anti-inflammatory, anti-oxidative, and anti-coagulant properties of CDP-

CHO (43, 52, 53), supports the therapeutic potential of this compound in the treatment or prevention of ARDS secondary to influenza infection. Perhaps more urgently, CDP-CHO also holds potential for treating COVID-19, which is associated with the development of

ARDS and coagulopathy (15, 54).

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4.6 Figures

Figure 4.1: Daily CDP-choline treatment following IAV infection rescues ATII cell phosphatidylcholine (PC) synthesis.

A) Diagram of CDP-choline (CDP-CHO) treatment regimen. B) Quantification of PC in BALF collected at 6 dpi from IAV-infected mice (and IAV-infected mice treated daily with CDP-CHO (n= 6-11 per group, *: p<0.05). C) Quantification of ATII cell choline and choline-phosphate (CHOLINE-P) at 6 dpi in mock-infected IAV-infected and IAV-infected mice treated daily with CDP-CHO and CDP-diacylgycerol (CHO/DAG-Tx,) (n=5-6 per group, **: p<0.005). D) Quantification of ATII cell PC at 6 dpi in mock-infected, IAV- infected, and IAV-infected mice treated daily with CDP-CHO and CDP-diacylgycerol (CHO/DAG-Tx) (n=5-6 per group, **: p<0.005). Data are presented as mean ± SEM.

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Figure 4.2: Daily CDP-choline treatment following IAV infection improves ATII cell mitochondrial physiology.

A-C) representative TEM images of ATII cells in the lungs of mock-infected (A), IAV- infected (B), and CDP-choline treated IAV-infected (C) mice at 6 dpi. Insets highlight mitochondria, denoted with white arrows. D) Mean density ratio of ATII cell tafazzin protein expression by western blot, compared to β-actin expression, in mock-, IAV-, and CDP-CHO treated IAV-infected mice at 6 dpi (n=3 mice per group, *: p=0.03). E) Representative western blot membrane evaluating tafazzin and β-actin protein expression in ATII cells isolated from mock-, IAV-, and CDP-CHO treated IAV-infected mice at 6 dpi. Data are presented as mean ± SEM.

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Figure 4.3: Daily CDP-choline treatment improves ATII cell mitochondrial function.

A) DiIC1(5) mean channel fluorescence (MCF) in ATII cells from mock-infected (blue), IAV-infected (red), and CDP-choline treated IAV-infected (purple) mice at 6 dpi (n=6-7 mice per group, *: p<0.05). B) Representative measurements of ATII cell oxygen consumption rate (OCR) over the timecourse of a Mito Stress Test assay for ATII cells from a mock (blue), IAV (red) infected, IAV-infected and CDP-choline treated (purple) mouse. C) Quantification of mean ATII cell OCR before the addition of inhibitors (BASAL) and following the addition of oligomycin (+OLIGO) and FCCP (+FCCP) (n>6 per group, *: p<0.05, #: p<0.01 **: p<0.005). D) Mean density ratio of citrate synthase expression, compared to β-actin expression, by western blot of ATII cells isolated from mock-, IAV-, and CDP-CHO treated IAV-infected mice at 6 dpi (n=3 per group, *: p=0.049. E) Representative western blot membrane evaluating citrate synthase and β- actin protein expression in ATII cells isolated from mock-, IAV-, and CDP-CHO treated IAV-infected mice at 6 dpi. Data are presented as mean ± SEM.

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Figure 4.4: CDP-choline treatment following IAV infection attenuates ARDS at 6 dpi.

A) Carotid arterial O2 saturation (SaO2) for IAV-infected mice (UNTx) and IAV-infected mice treated daily with CDP-choline (CDP-CHO) (n≥14 per group, #: p<0.001). B) Alveolar fluid clearance rate (AFC30 %) in mock-infected, IAV-infected, and IAV-infected CDP-CHO treated mice (n≥7 per group, #: p<0.001). C) Quantification of alveolar macrophages (AMs) and neutrophils (PMNs) in BALF from mock-infected (MOCK DAY 6), IAV-infected (FLU DAY 6), and CDP-CHO treated IAV-infected (FLU DAY 6 CDP- CHO) mice. D and E) Representative parenchymal histopathology in hematoxylin/eosin- stained lung tissue from IAV-infected (UNTx DAY 6) and CDP-CHO-treated IAV-infected (CDP-CHO-Tx DAY 6) mice.

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30. Aeffner F, Bratasz A, Flaño E, Powell KA, Davis IC. Post-infection A77-1726 treatment improves cardiopulmonary function in H1N1 influenza-infected mice. Am J

Respir Cell Mol Biol. 2012;47(4):543-51.

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Chapter 5. Future Directions

5.1 Phospholipid Synthesis

Although we and others have shown that IAV infection alters phospholipid synthesis in vivo (1-4), little is known about the mechanisms behind these changes. The majority of this work has been focused on the generation of pro-inflammatory lipid species during IAV infection; we are the first to assess the impact of altered phospholipid synthesis on cellular function. It remains unclear whether global changes in phospholipid metabolism are directly mediated by IAV or by the host response, and it is likely that both contribute. IAV derives its envelope from the host cell membrane, budding at sphingolipid-cholesterol rafts (5). As levels of both sphingomyelin and cholesterol are significantly increased in ATII cells from IAV-infected mice (4), it is possible that IAV upregulates production of these lipids to support budding, at the expense of other lipid species such as phospholipids. However, disruption of cholesterol synthesis in IAV- infected MDCK cells increased viral particle release (6), so it is also possible that host cells alter lipid synthesis as a way to reduce budding and viral spread.

Changes in phospholipid levels have also been reported in patients with ARDS, suggesting that this occurrence is not specific to viral infection. These studies have primarily been focused on pulmonary surfactant (7-10), as reduced lung compliance is a recognized component of ARDS (11, 12). However, these studies did not examine phospholipid levels within ATII cells, which synthesize and secrete pulmonary surfactant, or even in whole lung tissue. Defects in ATII cell phosphatidylcholine levels have been noted following other stimuli. Mice exposed to cigarette smoke for a short period of time

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had significantly reduced levels of phosphatidylcholine in ATII cells (13).

Phosphatidylcholine levels were also significantly reduced in ATII cells isolated from mice following in vitro exposure to acrolein, a pollutant that induces cellular oxidation

(14). Finally, halothane, an inhalation anesthetic, reduced phosphatidylcholine levels in

ATII cells isolated from rats (15). These reports indicate that changes in ATII cell phospholipid synthesis are not limited to infectious stimuli but can be induced by a number of harmful substances, suggesting a more generalized mechanism that alters phospholipid synthesis following ATII cell damage.

We have shown for the first time that changes in ATII cell PC synthesis are linked to reduced levels of its immediate precursor, CDP-choline, which is virtually undetectable in ATII cells following IAV infection. Why CDP-choline levels are so profoundly disrupted by IAV infection is unclear; the gene and protein expression of the enzyme that synthesizes CDP-choline, CTP:phosphocholine cytidylyltransferase (CCT-

α) is not altered in ATII cells following infection (unpublished data). Even basic mechanisms of CCT-α regulation remain uncertain. It is known that CCT-α is an amphitropic protein, which is activated when it binds to cellular membranes in loosely packed regions enriched for phosphatidic acid, diacylglycerol (DAG), and fatty acids

(16). ATII cells from IAV-infected mice contain significantly increased levels of DAG (4), yet still have a loss of CDP-choline, suggesting that there are additional levels of regulation for CCT-α activity. CCT-α is highly phosphorylated in a serine-proline enriched region at the C-terminal end, and dephosphorylation has been shown to induce

CCT-α membrane translocation, and therefore enzyme activity, however, although not in

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every context (16). A third hypothesis suggests that CCT-α activity is regulated by cellular localization. In many cell types, CCT-α is activated by translocation from the cytoplasm to the nucleus (16). Nuclear uptake of CCT-α is mediated by importin-α- importin-β, and loss of importin-β function ablated Kennedy pathway PC synthesis in yeast (17). Interestingly, IAV also uses importin-α/importin-β to enter the nucleus, and it has been reported that tumor necrosis factor alpha (TNF-α) induction by highly pathogenic avian IAVs (HPAIVs) reduces importin-α gene expression as an anti-viral mechanism in murine embryonic fibroblasts (MEFs) (18). It is possible that host cell downregulation of the importin-α-importin-β pathway following IAV infection also disrupts

CCT-α activity, reducing PC synthesis. Future work in our model of IAV-induced ARDS will examine these potential regulatory mechanisms to determine the mechanism of disrupted PC synthesis in ATII cells from infected mice.

5.2 Fatty Acid Metabolism

ATII cells have high mitochondrial content, three times more than other lung cell types (19), in order to fuel their many critical functions. It has long been thought that ATII cells rely largely on aerobic glycolysis and lactate to feed into mitochondrial energy production (20-22). Under this paradigm, ATII cells predominately use fatty acids to generate phospholipids for pulmonary surfactant, and will only derive energy from the breakdown of fatty acids under conditions of cellular stress (13). However, it was recently reported that ATII cells conduct β-oxidation of fatty acids to support energy production during homeostasis, and that LPS-induced acute lung injury disrupts ATII cell fatty acid oxidation and induces ATII cell dysfunction (23). Fatty acid metabolism

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connects two critical ATII cell processes- phospholipid synthesis for pulmonary surfactant production, and energy production to support ATII cell functions. Future work should therefore attempt to elucidate the role of fatty acid oxidation in ATII cell energy production in vivo during both homeostasis and various disease states.

5.3 Mitochondrial Dysfunction

We have reported defects in energy production, a major mitochondrial function, in ATII cells following lethal IAV infection. However, mitochondria have multiple other functions that were not extensively explored due to time and methodology limitations.

Other mitochondrial activities include roles in innate immune signaling (24, 25), calcium homeostasis (26-28), and initiation of apoptosis (29, 30). Furthermore, we were also limited in our exploration of ATII cell mitochondrial quality control (fusion, fission, and mitophagy) and mtDNA damage during IAV infection. Additionally, several IAV proteins have been proposed to interact with mitochondria (31-35). Recent reports have identified a myriad of interactions between IAV and proteins involved in intrinsic apoptotic signaling, which is mediated by mitochondria (36-39). Future research should be aimed at developing a more encompassing portrait of changes in ATII cell mitochondrial structure and functions following IAV infection, and assessing the extent to which these changes are directly induced by IAV infection versus the host response to the virus.

Mitochondrial dysfunction in other types of lung disease, especially chronic diseases such as idiopathic pulmonary fibrosis (IPF) and chronic obstructive pulmonary disorder (COPD), is increasingly recognized as a driver of disease (20, 40-46). Less is known about changes in mitochondrial function during ARDS, although mitochondrial

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dysfunction is increasingly recognized as an underlying cause of epithelial barrier damage (47). Our work shows that ATII cells experience a bioenergetic crisis during

IAV-induced ARDS, and that treatment with a therapeutic that can improve mitochondrial energy production attenuates ARDS in mice. ARDS is a heterogenous condition with many causes, and future assessment of ATII cell mitochondria in models of sterile ARDS would provide important information about the applicability of our work outside of IAV- induced ARDS. If a similar pattern of mitochondrial dysfunction and bioenergetic crisis occurs in ATII cells in other models of ARDS, it would reinforce not only that mitochondrial dysfunction is a component of this disease, but also that ATII cells play a central role in the development of ARDS.

Recently, the identification of two subtypes of ARDS- reactive and uninflamed- with substantially different mortality rates, has provided additional insight into the pathogenesis of this condition (48, 49). The reactive phenotype is associated with increased oxidative phosphorylation and mitochondrial dysfunction in blood leukocytes

(50). Future research into the behavior of ATII cell mitochondria in each subtype would help to further develop the characteristics of each subtype and identify mechanisms of

ARDS development that are common to all patients.

5.4 CDP-choline

Modulating mitochondrial function to prevent or treat ARDS is an attractive therapeutic strategy because it has the potential to attenuate disease with minimal off- target effects such as the development of anti-viral or antibiotic resistance, or systemic toxicity. We have shown that such a strategy is effective in mice by attenuating clinical

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signs of ARDS with CDP-choline, which restores mitochondrial energy production. Other animal models have also shown that CDP-choline is safe and effective in the treatment of critical illness (51, 52). Additionally, CDP-choline has an excellent safety profile in clinical studies (53). Future work in this area should assess the efficacy of CDP-choline for treating ARDS in larger animal models, and eventually human subjects. Additionally, given the dramatic restoration of ATII cell function following CDP-choline activity, this treatment strategy should be explored in other diseases when ATII cells are known to play a central role, such as COPD and IPF.

5.5 Final Thoughts

We have shown that lethal IAV infection induces dramatic changes in ATII cell mitochondrial structure and function in mice, without inducing intrinsic apoptosis. These changes include alterations in the phospholipid composition of mitochondrial membranes, altered expression of proteins that mediate mitochondrial biogenesis and fusion, altered electron transport chain function, and a global reduction in mitochondrial energy production in ATII cells following IAV infection. These mitochondrial defects are fundamentally linked to observed changes in ATII cell phospholipid synthesis in infected mice, as treatment with CDP-choline, a phospholipid precursor, rescues Kennedy pathway synthesis of the major mitochondrial phospholipid phosphatidylcholine. CDP- choline treatment simultaneously improves mitochondrial function and energy production in ATII cells while also attenuating IAV-induced ARDS in mice. This suggests that mitochondrial dysfunction plays a critical role in ARDS pathogenesis, which can be targeted with CDP-choline. Based on these promising results in mice, it is our hope that

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CDP-choline will prove to be an effective treatment for ARDS patients, reducing the impact of devastating viral diseases like influenza and COVID-19. We will continue to pursue this goal through further research on the mechanisms underlying mitochondrial and ATII cell dysfunction in IAV-induced ARDS.

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