Bishop Lab

Ewing sarcoma is treated primarily with combinations of chemotherapy drugs introduced decades ago. Although these treatments can be remarkably effective, we still do not fully understand why Ewing sarcoma is sensitive to these agents, why treatment fails in some patients or how tumors become resistant.

Our goal is to understand how the EWS::FLI1 fusion oncogene alters transcription, DNA repair and genome stability, and how Ewing sarcoma cells adapt to the resulting cellular stress. By defining the biology that allows these cancer cells to survive, we aim to identify vulnerabilities that can explain responses to existing therapies and provide opportunities for developing more effective, more targeted and less toxic treatments.

Our Ewing sarcoma program began by asking a fundamental question: why does chemotherapy work? Etoposide and other cytotoxic agents can be highly effective against Ewing sarcoma, yet the biological basis for this sensitivity was poorly understood. By studying how Ewing sarcoma cells respond to and survive etoposide-induced damage, our work led us to defects in homologous recombination and BRCA1 function, and ultimately to the discovery that the EWS::FLI1 fusion oncogene profoundly disrupts RNA polymerase II regulation, promotes R-loop accumulation and interferes with DNA repair.

These discoveries changed the way we think about Ewing sarcoma. EWS::FLI1 is best known as an abnormal transcription factor that rewires gene expression, but our work suggests that it also creates a fundamentally abnormal state of transcription. Ewing sarcoma cells exhibit altered RNA polymerase II phosphorylation and promoter-proximal pausing, increased transcription, accumulation of RNA:DNA structures called R-loops and associated replication and genome stress. Remarkably, the cells tolerate this state sufficiently well to survive and proliferate.

This has led to a major question that now drives our research: how do Ewing sarcoma cells survive the transcriptional stress created by their own oncogene, and does this adaptation create vulnerabilities that we can exploit therapeutically?

We approach this question at several levels. We investigate how EWS::FLI1 changes RNA polymerase II activity, including promoter-proximal pausing and the dynamics of transcription at individual genes. We study the RNA-processing and R-loop-regulating machinery that allows cells to manage abnormal transcription, including proteins such as DHX9 and pathways that regulate its function. We also examine how transcriptional stress intersects with DNA replication, BRCA1-dependent DNA repair and genome stability. Together, these studies seek to distinguish the abnormalities caused directly by EWS::FLI1 from the compensatory mechanisms that Ewing sarcoma cells have become dependent upon for survival.

Our team combines molecular and cell biology with functional genetic screens, CRISPR-based approaches, live-cell imaging, proteomics and genome-wide methods for measuring transcription, chromatin occupancy, R-loops, and DNA replication. We are also developing new experimental approaches that allow us to observe transcriptional behavior and R-loop biology dynamically and at increasingly high resolution.

Ultimately, our goal is to bring this biology back to the problem that motivated the work. We want to understand why Ewing sarcoma responds to particular therapies, why those treatments sometimes fail and what dependencies emerge as a consequence of the tumor’s unusual biology. Rather than identifying treatments solely because they kill Ewing sarcoma cells in experimental models, we aim to establish the biological reason for that sensitivity. This mechanistic understanding can provide a foundation for developing therapies that are more targeted, more effective and less toxic, as well as rational strategies for treating tumors that fail or become resistant to current therapy.

Our Ewing sarcoma research has received sustained support from the National Institutes of Health (NIH), Cancer Prevention and Research Institute of Texas (CPRIT), Stand Up To Cancer-Cancer Research UK, foundations and other organizations. Major current and previous awards supporting this research program include:

• NIH/NCI R01CA241554, “Dysregulated Transcription Processes in Ewing Sarcoma” (PI: Bishop, 2020–2026). Investigating the consequences of dysregulated transcription in Ewing sarcoma and the mechanisms that allow these cells to tolerate transcription-associated stress.
• NIH/NIGMS R01GM140127, “An Integrated Biochemical and Structural Approach to Delineating the Biology of EWSR1” (PI: Libich; Bishop, Co-Investigator, 2021–2026). Defining structure-function relationships of EWSR1.
• Jeff Gordon Foundation Award (PI: Bishop, 2024–2026). Investigating combinations of PARP inhibitors and ferroptosis-inducing agents as therapeutic strategies for Ewing sarcoma.
• Stand Up To Cancer-Cancer Research UK Pediatric Cancer New Discoveries Challenge, SU2C-CRUK RT6187, “Targeting R-loop Stability in Ewing Sarcoma” (PI: Bishop, 2021–2023). Investigating whether mechanisms that allow Ewing sarcoma cells to tolerate R-loops can be therapeutically targeted.
• CPRIT Individual Investigator Research Award RP150445, “Ewing Sarcoma, a Homologous Recombination Defective Disease” (PI: Bishop, 2015–2019). Defining the homologous recombination defect in Ewing sarcoma and exploiting this biology therapeutically.
• NIH/NCI R01CA152063, “Improving Etoposide Treatment of Ewing’s Sarcoma” (PI: Bishop, 2011–2018). Identifying mechanisms that allow Ewing sarcoma cells to survive etoposide and determining whether these pathways can be targeted to improve treatment.
• Andrew McDonough B+ Foundation Childhood Cancer Research Grant 614252, “Targeting RNA Processing Defects of Ewing Sarcoma” (PI: Bishop, 2019–2021). Investigating RNA-processing dependencies and their potential as therapeutic vulnerabilities.
• Rutledge Foundation Award, “Comparing the Efficacy of Olaparib and Talazoparib to Augment Ferroptosis Targeting of Ewing Sarcoma” (PI: Bishop, 2023–2025).

The program has additionally supported multiple NIH, Department of Defense, CPRIT, AACR-AstraZeneca and institutional predoctoral and postdoctoral fellowships investigating Ewing sarcoma transcription, R-loops, RNA processing, replication stress, DNA repair and therapeutic response.

R-loops are three-stranded nucleic acid structures that form when newly produced RNA remains paired with DNA. They are normal intermediates in many cellular processes, but when formed in the wrong place, at the wrong time or allowed to persist, they can interfere with transcription and DNA replication and threaten genome stability.

Our goal is to understand what distinguishes physiological from harmful R-loops, how cells regulate their formation and resolution and how R-loops interact with transcription, DNA repair, replication, and three-dimensional genome organization. We are also developing new experimental and computational tools to overcome longstanding limitations in R-loop detection, with the ultimate goal of identifying disease-associated R-loop dependencies that can be therapeutically targeted.

R-loops were once viewed primarily as unusual byproducts of transcription. It is now clear that they are widespread structures with important functions in transcription, chromatin organization, DNA repair and other aspects of genome regulation. At the same time, excessive or persistent R-loops can interfere with DNA replication, generate DNA damage and promote genome instability. A central challenge is therefore understanding when an R-loop is functional and when it becomes pathological.

Our interest in R-loops initially emerged from our studies of Ewing sarcoma. We discovered that the EWS::FLI1 fusion oncogene alters RNA polymerase II activity and causes widespread R-loop accumulation. Those findings led us to investigate more fundamental questions about how R-loops form, how cells resolve or tolerate them and how transcription, RNA processing, DNA replication and DNA repair collectively determine their biological consequences.

A major obstacle in answering these questions has been the technology available to study R-loops. Existing approaches often use different reagents for microscopy, genomic mapping and biochemical analysis, making results difficult to compare. Genome-wide R-loop measurements also typically require large numbers of cells, which has limited studies of rare cell populations and has made true single-cell analysis particularly challenging.

To address these limitations, we are developing a modular, genetically encoded R-loop sensor that can be temporally controlled and used across multiple experimental platforms. The same reagent can support live-cell and fixed-cell imaging, genomic mapping, affinity purification and potentially proteomic analyses, allowing us to examine the same underlying R-loop population in different ways. A major objective is to miniaturize R-loop detection toward low-input and ultimately single-cell genomic applications, which would allow us to investigate R-loop dynamics and heterogeneity that are hidden by population-level measurements.

In parallel, we develop computational approaches for analyzing R-loop genomic data. Our RLPipes and RLBase resources are being modernized to incorporate newer datasets, analytical approaches and telomere-to-telomere genome assemblies. Combining these computational resources with improved experimental detection should allow more rigorous comparisons across R-loop mapping technologies and biological systems.

We are also investigating how R-loops relate to the three-dimensional organization of the genome. Current studies examine how disruption of STAG2, a component of the cohesin complex frequently altered in cancer, changes chromatin architecture and how those changes influence transcription and R-loop formation. More broadly, we are interested in integrating R-loop measurements with chromatin architecture and functional genetic approaches to determine how genome organization influences where R-loops form and what consequences they have.

Ultimately, we want to move beyond simply cataloguing R-loops. By understanding where they arise, why they persist, how individual cells differ in their R-loop burden and what cellular mechanisms become necessary to tolerate them, we hope to identify fundamental principles of genome regulation as well as vulnerabilities created by pathological R-loop accumulation in cancer.

This work has developed through several interconnected awards supporting R-loop biology, transcription, computational tool development and trainee-led research. I would probably list the major ones as:

• NIH/NCI R01CA241554, “Dysregulated Transcription Processes in Ewing Sarcoma” (PI: Bishop, 2020–2026). Supports studies of transcriptional dysregulation, RNA processing, R-loops and associated vulnerabilities.
• Stand Up To Cancer-Cancer Research UK Pediatric Cancer New Discoveries Challenge, SU2C-CRUK RT6187, “Targeting R-loop Stability in Ewing Sarcoma” (PI: Bishop, 2021–2023). Investigated mechanisms that regulate R-loop stability and whether these dependencies can be therapeutically exploited.
• Mays Cancer Center PPG Development Award, “R-loop Biogenesis and Resolution by DNA Repair” (PI: Bishop, 2022–2024). Supported development of a collaborative program investigating fundamental mechanisms of R-loop formation and resolution.
• Reglagene Collaborative Research Grant, “The Clinical Potential and Application of Reglagene Compounds in R-loops” (PI: Bishop, 2021–2023). Investigated compounds that alter R-loop abundance and their effects in cells with different R-loop states.
• NIH/NIA F31AG072902, “The Dynamics and Impact of R-loops in Epigenetic Stability and Aging” (Henry Miller; Bishop, Mentor, 2021–2024).
• Department of Defense PRCRP Horizon Award, “Transcription, R-loops and RNA Splicing in Ewing Sarcoma” (Liesl Lawrence; Bishop, Mentor, 2019–2022).
• CPRIT Training Grant Fellowship, “Dedifferentiation in Ewing Sarcoma Mediated by R-loops Preserves Genomic Integrity” (Pramiti Mukhopadhyay; Bishop, Mentor, 2020–2021).

BRCA1 is best known for its role in repairing damaged DNA, but its functions extend well beyond DNA repair. Our goal is to understand how BRCA1 coordinates cellular responses to stress and contributes to normal mammary gland development, and how disruption of these functions influences both cancer development and therapeutic response.

A major focus of our work is the relationship between BRCA1 and NRF2, a central regulator of antioxidant and metabolic stress responses. We are investigating how BRCA1 controls NRF2 activity, how this relationship influences mammary differentiation and susceptibility to ferroptosis and whether these mechanisms can ultimately be leveraged to prevent or treat BRCA1-associated cancers.

Inherited mutations in BRCA1 dramatically increase the risk of developing breast and ovarian cancer. BRCA1-deficient breast tumors are frequently triple-negative breast cancers, an aggressive form of breast cancer with limited targeted treatment options. BRCA1’s best-characterized function is in homologous recombination, a pathway that accurately repairs damaged DNA. Loss of this repair function creates genomic instability but also provides a therapeutic vulnerability that can be exploited with treatments such as PARP inhibitors.

Our laboratory has studied BRCA1 and homologous recombination for more than two decades. Increasingly, however, our work has focused on functions of BRCA1 that extend beyond DNA repair. We are particularly interested in how BRCA1 coordinates different cellular responses to stress and how disruption of these functions contributes to both the development and treatment of cancer.

One major focus is the relationship between BRCA1 and NRF2, a transcription factor that coordinates antioxidant, metabolic, and other cellular stress responses. Our work indicates that BRCA1 influences NRF2 stability following cellular stress. We are now investigating the molecular basis of this relationship, including how BRCA1/BARD1 interacts with NRF2 and controls how long NRF2 persists after stress.

This interaction may have important consequences for ferroptosis, a form of regulated cell death caused by uncontrolled lipid oxidation. NRF2 controls several mechanisms that protect cells from ferroptosis, including glutathione metabolism and GPX4 activity. Our studies suggest that BRCA1 deficiency may weaken these defenses, creating an opportunity to selectively increase ferroptosis in BRCA1-deficient cancer cells. We are investigating whether ferroptosis induction can be rationally combined with PARP inhibitors and other therapies to improve treatment of BRCA1-deficient and triple-negative breast cancers.

Effective cancer therapy must also be considered in the context of the immune system. Natural killer (NK) cells recognize and destroy cancer cells and are increasingly being developed as cancer therapies. We are therefore investigating how PARP inhibition, ferroptosis induction and manipulation of NRF2 affect both tumor cells and NK cells. Our goal is to identify therapeutic conditions that preferentially kill cancer cells while preserving, or potentially enhancing, anti-tumor immune activity.

A parallel component of our research asks what BRCA1 does in the normal mammary gland and how disruption of those functions might contribute to cancer development. Our studies indicate that BRCA1 and NRF2 influence mammary epithelial differentiation, particularly during the extensive developmental changes that accompany pregnancy and lactation. We are investigating whether restoring NRF2-dependent differentiation can counteract consequences of BRCA1 loss and whether changing mammary epithelial cell state can ultimately alter the development or timing of BRCA1-associated tumors. This work provides a complementary opportunity to consider whether understanding BRCA1 biology could eventually inform approaches to cancer prevention as well as treatment.

We are also interested more broadly in how BRCA1 is allocated among its different cellular functions. BRCA1 participates in DNA repair, transcription, R-loop regulation and cellular stress responses. Our recent work has investigated how phosphorylation of the RNA polymerase II C-terminal domain controls recruitment of BRCA1 to transcriptional complexes. These studies support a direct interaction between the BRCA1 BRCT domain and Ser5-phosphorylated RNA polymerase II and suggest that cellular stress can alter this association, providing a potential mechanism for dynamically redistributing BRCA1 between transcription-associated and DNA damage-response functions.

Together, these studies examine how functions of BRCA1 beyond homologous recombination influence both the development of BRCA1-associated breast cancer and the response of established tumors to therapy. By defining how BRCA1 regulates mammary differentiation, NRF2-dependent stress responses, and susceptibility to ferroptosis, we aim to identify biological mechanisms that could ultimately be leveraged for cancer prevention or treatment.

This research builds on more than two decades of support for our studies of BRCA1, homologous recombination, DNA damage responses, cellular stress and therapeutic response. Relevant awards include:

• Basser Center for BRCA External Grant Program, “Identifying the Role of BRCA1 in the Mammary Gland Thus Leading to Therapeutic Targets” (PI: Bishop, 2021–2023; $100,000). This project investigated the normal function of BRCA1 during mammary gland development and provided an important foundation for our current studies of BRCA1, NRF2, mammary differentiation and breast cancer susceptibility.
• NIH/NIEHS R15ES019128, “Comparing and Contrasting the Biology of Damage Survival at a Genomic Level” (PI: Bishop, 2010–2013; $300,000 direct costs). This work used functional genomic approaches to identify conserved genes and pathways that determine cellular survival following different forms of DNA damage.
• NIH/NIEHS K22ES012264, “A Screen for Damage Response Using Recombination” (PI: Bishop, 2004–2010; $324,000). This award supported genome-scale studies of genes regulating homologous recombination following DNA damage and helped establish the laboratory’s long-standing program in DNA repair and genome stability.
• Max and Minnie Tomerlin Voelcker Fund Young Investigator Award, “Determining Cisplatin Survival Factors to Augment Ovarian Cancer Treatment” (PI: Bishop, 2010–2015; $409,092 direct costs). This work sought to identify mechanisms that allow ovarian cancer cells to survive cisplatin and determine whether those mechanisms could be targeted to improve therapy.
• CPRIT Training Grant Fellowship, “Estrogen-Induced R-loops are NRF2 and BRCA1 Dependent” (Carolina Romero; Bishop, Mentor, 2015–2016). This trainee-led project investigated connections among BRCA1, NRF2, estrogen-induced transcription and R-loop biology.
• DoD Breast Cancer Research Program Postdoctoral Fellowship W81XWH-14-1-0026, “14-3-3sigma Mislocalization in Metastatic and Chemoresistant Breast Cancer” (Alfeu Zanotto-Filho; Bishop, Supervisor, 2014). This project investigated mechanisms contributing to metastatic and chemotherapy-resistant breast cancer.

Current studies extend this foundation to define the molecular relationship between BRCA1/BARD1 and NRF2, determine how this pathway controls ferroptosis and therapeutic response, investigate its role in mammary differentiation and cancer development and identify treatment combinations that can target BRCA1-deficient cancer while preserving anti-tumor immune activity.

Ataxia-telangiectasia (A-T) is a rare inherited disorder caused by loss of the ATM protein and characterized by progressive neurological disease, immune dysfunction, cancer predisposition and metabolic and vascular abnormalities. Although ATM has been studied extensively for its role in responding to DNA damage, these functions do not adequately explain many of the progressive manifestations of A-T.

Our goal is to identify the underlying cellular and metabolic consequences of ATM loss that contribute to disease and determine whether these defects can be bypassed or corrected therapeutically. Our recent work has uncovered a role for ATM in regulating amino acid transport and metabolism, providing both a new framework for understanding A-T and potential opportunities for intervention.

Our laboratory has a longstanding interest in ATM biology and ataxia-telangiectasia (A-T), beginning with early studies of how ATM regulates homologous recombination and genome stability. More recently, we returned to a fundamental question: why does loss of a DNA damage-response protein produce the remarkably diverse and progressive features of A-T, including neurodegeneration, telangiectasia, immune dysfunction, cancer predisposition and metabolic disease?

Our recent work identified an unexpected connection between ATM and amino acid transport. We discovered that ATM phosphorylates CD98HC, a protein that partners with several different amino acid transporters and helps direct them to the cell surface. ATM-dependent phosphorylation accelerates movement of newly synthesized CD98HC-containing transporters to the cell membrane, allowing cells to rapidly adjust amino acid transport in response to stress. When ATM is absent, this response is impaired.

One consequence is altered transport of cystine and glutamate. ATM-deficient cells import less cystine and are less able to export glutamate, resulting in decreased glutathione and accumulation of intracellular glutamate. Importantly, our studies indicate that this is not simply a consequence of increased oxidative stress. Instead, it reflects a specific metabolic defect resulting from impaired regulation of amino acid transport.

Because CD98HC partners with multiple amino acid transporters, this mechanism has the potential to affect different tissues in different ways. In endothelial cells, for example, loss of ATM also impairs transport of arginine, an amino acid required for nitric oxide production and normal blood-vessel formation. Bypassing the affected transport systems with cell-permeable sources of cysteine and arginine restored aspects of normal endothelial function, providing a potential mechanistic connection to the vascular abnormalities characteristic of A-T.

We found related consequences in pancreatic cells and ATM-deficient mice. Pancreatic cells lacking normal ATM activity accumulate glutamate, have altered glutathione and mitochondrial metabolism, and show impaired hormone secretion. ATM-deficient mice progressively develop abnormal glucose regulation, loss of glucagon-producing pancreatic cells, glutamate and glutamine accumulation in pancreatic islets and hepatic lipid accumulation.

Importantly, understanding the mechanism suggested a potential intervention. N-acetylcysteine (NAC) can bypass the impaired cystine transport pathway by providing cells with an alternative source of cysteine. In ATM-deficient mice, NAC reduced glutamate accumulation in pancreatic islets, restored glucagon-positive cells and rescued abnormalities in glucose regulation and hepatic lipid accumulation. This provides an example of the approach that guides our laboratory more broadly: understanding the underlying biology of disease can reveal therapeutic opportunities that would otherwise be difficult to recognize.

An important emerging question is whether the same biology contributes to the progressive neurological manifestations of A-T. CD98HC associates with several amino acid transporters that have important functions in the nervous system, raising the possibility that loss of ATM could alter the availability or intracellular balance of amino acids required for normal neuronal function. Our published work therefore identified studies in human neurons and iPSC-derived neuronal models as an important direction for determining whether ATM-dependent regulation of amino acid transport contributes to neurological disease.

Our longer-term goal is to determine which manifestations of A-T can be traced to specific, actionable consequences of ATM loss and whether those defects can be bypassed or corrected therapeutically. Although A-T is currently a smaller and emerging component of our laboratory’s research program, it illustrates the same principle that guides our cancer research: define the underlying biology, understand how it produces disease and use that knowledge to identify rational opportunities for intervention.

Direct support for this work has included:

• Greehey Children’s Cancer Research Institute Exploratory Experimental Fund, “Metabolomics in ATM-Inhibited Cells” (PI: Bishop, 2019–2020; $10,000). This pilot award supported stable-isotope tracing studies to determine how loss of ATM activity alters glucose and glutamine metabolism. These experiments contributed to the development of our current model linking ATM loss to altered amino acid metabolism.
• NIH/NCI T32CA148724, Training Grant on the Biology of Cancer (Predoctoral Traineeship to Sonal Tonapi; Bishop, Mentor, 2013–2015; $60,460). This fellowship supported studies of oxidative stress responses, nucleotide metabolism, mutation and metabolic dysfunction associated with ATM loss.

Additional support from NIH, CPRIT, institutional and philanthropic sources contributed resources to the broader research program that ultimately led to our identification of the ATM-CD98HC amino acid transport pathway.