Guidelines for Evaluating Endothelial Function in Vascular Tissue.

Guidelines for Evaluating Endothelial Function in Vascular Tissue - PMC Skip to main content An official website of the United States government Here's how you know Here's how you know Official websites use .gov A .gov website belongs to an official government organization in the United States. Secure .gov websites use HTTPS A lock ( Lock Locked padlock icon ) or https:// means you've safely connected to the .gov website. Share sensitive information only on official, secure websites. Search Log in Dashboard Publications Account settings Log out Search… Search NCBI Primary site navigation Search Logged in as: Dashboard Publications Account settings Log in Search PMC Full-Text Archive Search in PMC Journal List User Guide PERMALINK Copy As a library, NLM provides access to scientific literature. Inclusion in an NLM database does not imply endorsement of, or agreement with, the contents by NLM or the National Institutes of Health. Learn more: PMC Disclaimer | PMC Copyright Notice Am J Physiol Heart Circ Physiol . Author manuscript; available in PMC: 2026 Apr 17. Published in final edited form as: Am J Physiol Heart Circ Physiol. 2026 Mar 9;330(5):H1600–H1672. doi: 10.1152/ajpheart.00656.2025 Search in PMC Search in PubMed View in NLM Catalog Add to search Guidelines for Evaluating Endothelial Function in Vascular Tissue Cameron G McCarthy Cameron G McCarthy 1. Department of Cell Biology and Anatomy, Cardiovascular Translational Research Center, University of South Carolina, School of Medicine, Columbia, SC, USA Find articles by Cameron G McCarthy 1 , Christian Aalkjær Christian Aalkjær 2. Department of Biomedicine, Aarhus University, Aarhus, Denmark Find articles by Christian Aalkjær 2 , Pooneh Bagher Pooneh Bagher 3. Department of Cellular and Integrative Physiology, University of Nebraska Medical Center, Omaha, NE, USA Find articles by Pooneh Bagher 3 , Andreas M Beyer Andreas M Beyer 4. Departments of Medicine and Physiology, Cardiovascular Research Center, at the Medical College of Wisconsin, Milwaukee, WI, USA. 5. A.I. Virtanen Institute for Molecular Sciences, Faculty of Health Sciences University of Eastern Finland Find articles by Andreas M Beyer 4, 5 , Ebbe Boedtkjer Ebbe Boedtkjer 2. Department of Biomedicine, Aarhus University, Aarhus, Denmark Find articles by Ebbe Boedtkjer 2 , Gisele F Bomfim Gisele F Bomfim 6. Health Education Research Center, Institute of Health Sciences, Federal University of Mato Grosso, Sinop, MT, Brazil Find articles by Gisele F Bomfim 6 , Jerome W Breslin Jerome W Breslin 7. Department of Molecular Pharmacology and Physiology, Morsani College of Medicine, University of South Florida, FL, USA Find articles by Jerome W Breslin 7 , Ana M Briones Ana M Briones 8. Department of Pharmacology, Faculty of Medicine, Universidad Autónoma de Madrid, Hospital La Paz Institute for Health Research (IdiPaz), Madrid, Spain; CIBER Cardiovascular, Spain Find articles by Ana M Briones 8 , Jorge A Castorena-Gonzalez Jorge A Castorena-Gonzalez 9. Dept. of Pharmacology, Tulane University, New Orleans, Louisiana, USA Find articles by Jorge A Castorena-Gonzalez 9 , Tiago J Costa Tiago J Costa 10. Department of Pharmacology, Institute of Biomedical Science, University of Sao Paulo, Brazil Find articles by Tiago J Costa 10 , Zhiyu Dai Zhiyu Dai 11. Department of Medicine, Division of Pulmonary & Critical Care Medicine, Washington University, St. Louis, MO, USA Find articles by Zhiyu Dai 11 , Ana P Davel Ana P Davel 12. Department of Structural and Functional Biology, Institute of Biology, Universidade Estadual de Campinas, Campinas, Brazil Find articles by Ana P Davel 12 , Scott Earley Scott Earley 13. Department of Pharmacology and Physiology, University of Rochester Medical Center, Rochester, NY Find articles by Scott Earley 13 , Julie K Freed Julie K Freed 14. Department of Anesthesiology, Department of Physiology, Cardiovascular Research Center, Medical College of Wisconsin, Milwaukee, WI, USA Find articles by Julie K Freed 14 , Christopher Garland Christopher Garland 15. Department of Pharmacology, University of Oxford, Oxford, UK Find articles by Christopher Garland 15 , Brant E Isakson Brant E Isakson 16. Molecular Physiology and Biological Physics, Robert M. Berne Cardiovascular Research Center, University of Virginia, Charlottesville, VA, USA Find articles by Brant E Isakson 16 , Thomas A Jepps Thomas A Jepps 17. Vascular Biology Group, University of Copenhagen, Copenhagen, Denmark Find articles by Thomas A Jepps 17 , Joanna Kalucka Joanna Kalucka 2. Department of Biomedicine, Aarhus University, Aarhus, Denmark Find articles by Joanna Kalucka 2 , Boris Lavanderos Boris Lavanderos 13. Department of Pharmacology and Physiology, University of Rochester Medical Center, Rochester, NY Find articles by Boris Lavanderos 13 , Ayako Makino Ayako Makino 18. The Herbert Wertheim UF Scripps Institute University of Florida, Jupiter, FL, USA Find articles by Ayako Makino 18 , Charles E Norton Charles E Norton 19. Department of Medical Pharmacology and Physiology, University of Missouri, Columbia, MO, USA Find articles by Charles E Norton 19 , Steven S Segal Steven S Segal 19. Department of Medical Pharmacology and Physiology, University of Missouri, Columbia, MO, USA Find articles by Steven S Segal 19 , Wenbin Tan Wenbin Tan 1. Department of Cell Biology and Anatomy, Cardiovascular Translational Research Center, University of South Carolina, School of Medicine, Columbia, SC, USA Find articles by Wenbin Tan 1 , Aaron J Trask Aaron J Trask 20. Center for Cardiovascular Research and the Heart Center, The Abigail Wexner Research Institute at Nationwide Children’s Hospital; Department of Pediatrics, College of Medicine, The Ohio State University Wexner Medical Center, Columbus, OH, USA Find articles by Aaron J Trask 20 , Calum Wilson Calum Wilson 21. Strathclyde Institute of Pharmacy and Biomedical Sciences, University of Strathclyde, Glasgow, UK. Find articles by Calum Wilson 21 , Scott D Zawieja Scott D Zawieja 19. Department of Medical Pharmacology and Physiology, University of Missouri, Columbia, MO, USA Find articles by Scott D Zawieja 19 , Camilla F Wenceslau Camilla F Wenceslau 1. Department of Cell Biology and Anatomy, Cardiovascular Translational Research Center, University of South Carolina, School of Medicine, Columbia, SC, USA Find articles by Camilla F Wenceslau 1, * Author information Article notes Copyright and License information 1. Department of Cell Biology and Anatomy, Cardiovascular Translational Research Center, University of South Carolina, School of Medicine, Columbia, SC, USA 2. Department of Biomedicine, Aarhus University, Aarhus, Denmark 3. Department of Cellular and Integrative Physiology, University of Nebraska Medical Center, Omaha, NE, USA 4. Departments of Medicine and Physiology, Cardiovascular Research Center, at the Medical College of Wisconsin, Milwaukee, WI, USA. 5. A.I. Virtanen Institute for Molecular Sciences, Faculty of Health Sciences University of Eastern Finland 6. Health Education Research Center, Institute of Health Sciences, Federal University of Mato Grosso, Sinop, MT, Brazil 7. Department of Molecular Pharmacology and Physiology, Morsani College of Medicine, University of South Florida, FL, USA 8. Department of Pharmacology, Faculty of Medicine, Universidad Autónoma de Madrid, Hospital La Paz Institute for Health Research (IdiPaz), Madrid, Spain; CIBER Cardiovascular, Spain 9. Dept. of Pharmacology, Tulane University, New Orleans, Louisiana, USA 10. Department of Pharmacology, Institute of Biomedical Science, University of Sao Paulo, Brazil 11. Department of Medicine, Division of Pulmonary & Critical Care Medicine, Washington University, St. Louis, MO, USA 12. Department of Structural and Functional Biology, Institute of Biology, Universidade Estadual de Campinas, Campinas, Brazil 13. Department of Pharmacology and Physiology, University of Rochester Medical Center, Rochester, NY 14. Department of Anesthesiology, Department of Physiology, Cardiovascular Research Center, Medical College of Wisconsin, Milwaukee, WI, USA 15. Department of Pharmacology, University of Oxford, Oxford, UK 16. Molecular Physiology and Biological Physics, Robert M. Berne Cardiovascular Research Center, University of Virginia, Charlottesville, VA, USA 17. Vascular Biology Group, University of Copenhagen, Copenhagen, Denmark 18. The Herbert Wertheim UF Scripps Institute University of Florida, Jupiter, FL, USA 19. Department of Medical Pharmacology and Physiology, University of Missouri, Columbia, MO, USA 20. Center for Cardiovascular Research and the Heart Center, The Abigail Wexner Research Institute at Nationwide Children’s Hospital; Department of Pediatrics, College of Medicine, The Ohio State University Wexner Medical Center, Columbus, OH, USA 21. Strathclyde Institute of Pharmacy and Biomedical Sciences, University of Strathclyde, Glasgow, UK. * Corresponding author: Dr. Camilla Ferreira Wenceslau, Department of Cell Biology and Anatomy, University of South Carolina School of Medicine, 6311 Garners Ferry Rd, Columbia, South Carolina, 29208 US, [email protected] Issue date 2026 May 1. PMC Copyright notice PMCID: PMC13085230  NIHMSID: NIHMS2157067  PMID: 41801061 The publisher's version of this article is available at Am J Physiol Heart Circ Physiol Abstract The endothelium plays a central role in maintaining vascular homeostasis by orchestrating vascular tone, inflammation, healing, permeability, and thrombosis. Assessing endothelial function in vascular tissue is essential for understanding the cellular and molecular mechanisms underlying cardiovascular physiology and pathology. Traditional approaches, such as wire and pressure myography, have been instrumental in defining endothelium-dependent responses and identifying key pharmacological targets. However, the complexity and heterogeneity of endothelial cells across vascular beds, and their dynamic phenotypic changes in health and disease, necessitate the incorporation of new investigative strategies. Emerging methodologies, including bulk and single-cell transcriptomics, proteomics, and advanced imaging, now provide unprecedented insights into endothelial cell diversity and function. A team of leading experts in the field, who collectively reached a consensus on the most widely used techniques to evaluate endothelial function, developed these guidelines. The document establishes best practices for assessing endothelial function, from endothelial cell cultures to isolated vascular tissues, integrating conventional functional assays with modern molecular approaches. By fostering methodological consistency and embracing innovation, our goal is to enhance rigor, reproducibility, understanding, and discovery in endothelial biology. Keywords: Endothelial Cells, Endothelium, Vessels, Vasodilation, Methods 1. INTRODUCTION Overview of the Consensus Guidelines In recent years, the scientific community has become increasingly aware of the challenges posed by inconsistent and irreproducible experimental findings. These issues often stem from variability in study design, lack of methodological standardization, and incomplete reporting of experimental details. To address these concerns, there has been a growing emphasis by funding agencies, leading journals such as American Journal of Physiology - Heart and Circulatory Physiology, and the broader research community on establishing clear guidelines that promote rigor, transparency, and reproducibility in experimental research. Adopting standardized best practices in commonly used techniques is essential to ensure that findings are robust, comparable across studies, and contribute meaningfully to the advancement of vascular science. This document was developed by a diverse team of investigators in vascular biology, representing multiple countries and continents, and spanning all academic stages, from early-career scientists to senior leaders in the field. Through collaborative discussions and shared expertise, this group reached a consensus on the most widely used and validated techniques to evaluate endothelial cell (EC) function in vascular tissue. By following these recommendations, researchers can foster methodological consistency, improve reproducibility, and advance the quality and translational impact of endothelial-focused vascular research. In these guidelines, we did not focus on how to mount arteries using wire or pressure myographs or on protocols for evaluating vascular contractility, topics that are comprehensively addressed in the Guidelines for Measurement of Vascular Function and Structure in Isolated Arteries and Veins ( 1 ). The scope of this consensus document is restricted to ex vivo assessments of endothelial function in isolated vascular tissues and complementary studies performed in EC culture systems. We recognize that endothelial function can also be evaluated in vivo in various animal models and in humans; however, these whole-organism approaches rely on different principles, experimental platforms, and interpretive frameworks. As such, they are not covered in the present guidelines, which focus specifically on standardized approaches for isolated arteries, microvascular segments, and cell-based endothelial assays. Instead, we focused on the evaluation of EC function in intact vascular tissue preparations and isolated EC systems. Here, we provide guidance on key aspects of experimental design that are specific to EC function, such as the choice and concentration of pre-constrictors for different vascular beds, when to initiate endothelium-dependent relaxation, mechanism of action of pharmacological agents, and interpretation of physiological and pharmacological responses (including flow-induced dilation, biphasic responses and endothelium-induced contraction in pathological conditions). We also discussed how tone maintenance can affect subsequent relaxation responses and data interpretation. Beyond myography-based assessments, this document covers the use of techniques such as EC Ca 2+ imaging in intact vessels, isolated microvascular endothelial tubes, EC metabolism and stiffness, barrier integrity assays, and other emerging molecular imaging, and electrophysiological approaches that capture EC heterogeneity and dynamic behavior. Finally, we leveraged the development of the present consensus guidelines as an opportunity to redefine the outdated concept of endothelial dysfunction, which is addressed in the Role of Endothelium in Diseases section and in the final section of this document under Perspectives . A Short History of Endothelial Cell Research Ever since Furchgott and Zawadzki published their seminal paper ( 2 ), which hopefully is included in every cardiovascular curriculum in the world, the endothelium has taken center stage in cardiovascular physiology. The paper not only opened our eyes to the importance of the endothelium, it forcefully demonstrated how important it is in science to think twice when we are confronted with an unexpected result. So, if it has been some time since you read it, please consider reading it again! In addition to this classic work, it is also worth reading Paul Vanhoutte’s account ( 3 ) of the pivotal meeting where both Robert Furchgott and Louis Ignarro informed the field that what was then called Endothelial Derived Relaxing Factor (EDRF) was, in fact, a gas, and nitric oxide (NO) was henceforth added to our vocabulary. Of course, we now appreciate that the collection of ECs in the body constitutes a major organ that, in addition to forming an important barrier between the blood and the tissue, generates a plethora of molecules while being responsive to many other molecules and to the flow of blood through the vessel lumen. The molecules are released predominantly in a paracrine and autocrine fashion to control a myriad of processes in the body including blood vessel diameter, the fluidity of the blood, barrier function of the endothelial monolayer, immunological function and transport, e.g. of oxygen, carbon dioxide, nutrients and metabolites. We also know now that pathology in the ECs can have dire consequences for the entire body, which is unsurprising given all of these key functions. Multiple roles for the endothelium are found not only in mammals, but also in reptiles, fish and birds, so nature has exploited the endothelium extensively throughout the animal kingdom! It follows that the favorite subject of many researchers is EC biology, and these guidelines are dedicated to considering how the endothelium can be studied in both a consistent and comprehensive manner. Basic Principles of Endothelial Function and Subpopulations of Endothelial Cells Endothelial-Mediated Vasodilation and Vasoconstriction Historically, we often think of the vasoactive function of the ECs in terms of vasodilation. This is justified by the tremendous physiological importance of endothelium-mediated relaxation of vascular smooth muscle cells (VSMCs) reflecting the synthesis and release of NO and prostacyclin and the generation of endothelial-derived hyperpolarization (EDH). Furthermore, the multiple vasodilator pathways add to the rich literature on endothelial-mediated vasodilation, giving them primacy in our minds. It is well known (i) which three enzymes are responsible for producing NO, (ii) how each is regulated through transcriptional and translational pathways, second messengers, and false substrates, and (iii) the complex metabolism of NO. For prostacyclin, we also know (i) the key enzymes involved in its production and (ii) the metabolic pathways responsible for its removal. The concept of EDH has also evolved over the last twenty or so years, and is now known to encompass not only coupling via diffusible chemical mediators, including K + , lipid mediators and hydrogen peroxide (H 2 O 2 ), but also direct electrical coupling via myoendothelial gap junctions, with research facilitating our understanding of the EC protrusions that form these contacts with the overlying VSMCs through the internal elastic lamina. It has also become apparent that these thin membrane protrusions contain an array of signaling complexes influencing vascular function. No matter how much we concentrate on understanding the vasodilator role of the endothelium, we should not forget that the endothelium also synthesizes and releases vasoconstrictor substances. Most well-known of these is endothelin, which has both autocrine and paracrine vasoconstrictor actions. Moreover, arachidonic acid metabolites and local formation of angiotensin II can also cause vasoconstriction and may have important roles, particularly in some vascular pathologies. A more complete understanding of endothelial-mediated vasoconstriction under different conditions is highly warranted. Endothelial Control of Blood Fluidity Extensive research has highlighted the role of the vascular endothelium in the development of atherosclerosis and thrombus formation. Several key vasodilatory substances, including NO and prostacyclin, also signal directly to circulating blood components, where they act to prevent platelet activation and inhibit clot formation. In fact, these antithrombotic actions may represent the primary physiological role of prostacyclin. Under certain conditions, however, endothelial cells can shift toward a pro-coagulant phenotype and produce factors such as thromboxane A 2 , tissue factor, and plasminogen activator inhibitor-1 (PAI-1). The endothelium therefore plays a critical role in maintaining the balance between preserving blood fluidity under normal conditions and enabling hemostasis when necessary. Unfortunately, this same system can contribute to pathological thrombus formation in the setting of advanced atherosclerotic lesions. The Role of the Endothelium in Barrier and Immune Function The endothelium serves as a critical barrier function, selectively regulating the exchange of molecules and cells between the blood and surrounding tissues. In certain organs, such as the brain, it forms a highly restrictive interface, known as the blood-brain barrier (BBB), while in others, like the kidney, liver, and spleen, it permits the passage of molecules or even cells between the bloodstream and parenchyma. This selective permeability is particularly relevant in inflammation, where ECs upregulate adhesion molecules (e.g., ICAM, VCAM) to facilitate leukocyte attachment and migration into underlying tissues, a process known as leukocyte extravasation. This immune response further amplifies inflammation, leading to increased endothelial permeability and the leakage of proteins from the blood into the parenchyma. Such mechanisms are central to the pathogenesis of cardiovascular diseases, such as atherosclerosis, hypertension and dementia, and contribute to inflammatory injuries throughout the body, including within the walls of small arteries, where emerging evidence suggests inflammatory involvement. The Heterogeneity of Endothelial Cells Recent advances in single-cell “OMICs” have uncovered remarkable ECs heterogeneity across different organs, reflecting their specialized roles in maintaining tissue homeostasis. The functional specialization of ECs is largely determined by their microenvironment and the specific needs of the organs in which they reside ( Fig. 1 ). Single-cell transcriptomic studies have been instrumental in uncovering the complexity of EC subtypes across different vascular beds, highlighting both organotypic and non-organotypic heterogeneity. Figure 1. Open in a new tab Endothelial cells (EC) exhibit organ-specific heterogeneity, adapting structurally and functionally to each tissue. Image taken with permission from ( 7 ). Organ-specific EC diversity is particularly evident, e.g. in the brain ( 4 ), where ECs exhibit gradual transcriptional changes along the arteriovenous axis, a process known as zonation. Zonation refers to the gradual, spatially organized changes in endothelial cell gene expression and function that occur along the arteriovenous axis within an organ. For example, in the brain, this means that arterial, capillary, and venous ECs display continuous, position-dependent transcriptional differences that support distinct physiological roles, such as barrier integrity, molecular transport, and metabolic regulation. This zonation is essential for the selective transport of molecules across the BBB, with arterial ECs enriched in transcription factors and capillary and venous ECs predominantly express transporter genes. A comparable pattern is seen in the liver sinusoids, where ECs are distributed along the portal vein-central vein axis, though it is worth noting that differences do exist between human and murine zonation profiles. Similar heterogeneity is observed in the lungs ( 5 ), where alveolar microvasculature consists of two distinct subtypes: (i) aerocytes and (ii) general capillary ECs (gCap). Aerocytes, characterized by their large, thin morphology, are closely associated with alveolar type I epithelial cells and express genes involved in adhesion and immune cell sequestration, optimizing them for gas exchange and leukocyte trafficking. In contrast, gCap cells are located in thicker regions of the pulmonary stroma, and their function acts in vasomotor regulation and alveolar capillary homeostasis, also serving as progenitor cells for vascular repair. The aorta, a major conductance artery, receives blood directly from the left ventricle and must withstand high transmural pressure. In response, its wall structurally adapts, and ECs are exposed to different shear stress patterns, high shear stress in the outer curvature and low shear stress in the inner curvature ( 6 )( 7 ) ( Fig. 2 ). This results in location-specific EC phenotypes. Specifically, a key recent study profiled single-cell transcriptional signatures in the mouse aorta and identified ten major cell clusters, including fibroblasts, vascular smooth muscle cells (VSMCs), ECs, and immune cells ( 6 ). Among ECs, three distinct subpopulations emerged. (i) EC1 expressed high levels of canonical EC markers like VCAM-1. (ii) EC2 showed elevated expression of lipid transport and angiogenesis-related genes, such as Cd36 (fatty acid translocase), Lpl (lipoprotein lipase), and Fms-like tyrosine kinase 1, also known as vascular endothelial growth factor receptor (VEGFR)-1 ( 6 ). (iii) EC3 was marked by lymphatic EC-associated genes, notably lymphatic vessel endothelial hyaluronan receptor 1 (LYVE-1). Spatial mapping revealed EC1 predominated in the inner curvature, while EC2 was enriched in the outer curvature, patterns likely driven by local shear stress, underscoring the link between transcriptional heterogeneity and regional endothelial specialization. These adaptations occur in various tissues influenced by pressure, flow, hormonal, and local cues ( 6 )( 7 ). Figure 2. Open in a new tab Distinct populations of endothelial cells (ECs) in the inner and outer curvatures of the aortic arch. ECs exhibit phenotypic differences in response to shear stress and blood flow patterns. Image reproduced with permission from reference ( 7 ). During development, EC heterogeneity emerges early, even before full vascular specialization occurs. In the developing heart, the endocardium, a specialized endothelial layer, not only serves as a barrier but also gives rise to different cardiac cell types. Heart valve formation begins with endocardial cushion cells undergoing endothelial-to-mesenchymal transition (EndMT), a process that is tightly regulated by surrounding myocardial signals. Early scRNA-seq studies of embryonic ECs have identified distinct subpopulations, suggesting that ECs differentiation occurs in a stepwise manner. The first vascular networks arise from mesodermal precursors through vasculogenesis, with primitive ECs initially appearing as non-specialized cells. However, scRNA-seq data from early mouse embryos indicate that EC identity is established earlier than previously believed, with specific transcriptional signatures distinguishing ECs based on their anatomical origin. ECs are highly plastic, capable of shifting their phenotype in response to both genetic and environmental cues ( 7 )( 8 )( 9 ). This plasticity is evident during embryonic development, where hemogenic ECs in the aorta-gonad-mesonephros region give rise to hematopoietic stem cells through endothelial-to-hematopoietic transition (EndHT), a process regulated by factors like runt-related transcription factor 1 and neurogenic locus notch homolog protein signaling. Interestingly, EndHT-like activity has also been observed in adult mice during tissue repair and inflammation, although its mechanisms remain under investigation. EndHT may be reactivated under pathological conditions, including disturbed blood flow. Beyond EndHT, ECs can undergo endothelial-to-immune-cell-like transition, particularly in response to proinflammatory stimuli and disturbed flow. Recent studies using scRNA-seq and scATAC-seq have identified ECs expressing immune markers such as complement C1q subcomponent subunit C and complement component 5a receptor 1, both in mouse aortic tissue and in vitro in human aortic ECs exposed to oscillatory shear stress ( 9 ). Emerging evidence also points to EC transdifferentiation into osteoblast-like cells, especially in vascular calcification and bone-related diseases. This transition is associated with regulators such as runt-related transcription factor, bone morphogenetic protein 2, and wingless-related integration site signaling, and has clinical relevance in atherosclerosis and diabetes, where increased numbers of osteocalcin-expressing endothelial progenitor cells correlate with vascular calcification ( 10 )( 11 )( 12 ). Among these transitions, EndMT is the most extensively studied ( 13 )( 14 ). In EndMT, ECs lose endothelial markers (e.g., VE-cadherin, CD31, so known as platelet endothelial cell adhesion molecule-1, PECAM-1) and acquire mesenchymal traits (e.g., alpha-smooth muscle actin, vimentin), driven by transcription factors such as snail family transcriptional repressor and zinc finger e-box–binding homeobox 1. This enhances their migratory capacity and extracellular matrix production, resembling invasive cell phenotypes. It is known that ECs switch phenotypes under adverse circumstances to restore their physiological functions. However, chronic dysregulation of this phenotypic switching could be associated with the development of cardiovascular diseases ( Fig. 3 ). Figure 3. Open in a new tab Endothelial cells (ECs) undergo phenotypic transitions that contribute to the development of cardiovascular disease. Image reproduced with permission from reference ( 7 ). Understanding EC diversity and phenotypic plasticity is crucial for advancing therapeutic strategies targeting vascular dysfunction. Insights into endothelial zonation could improve drug delivery across the BBB, while knowledge of sex- and age-related endothelial changes may inform personalized approaches for treating cardiovascular and neurodegenerative diseases. Likewise, a deeper understanding of EC development and differentiation could enhance regenerative medicine and vascular tissue engineering. As research continues to unravel the molecular mechanisms governing EC heterogeneity, it holds promise for refining disease models and developing targeted interventions for vascular pathologies. Role of Endothelium in Diseases Redefinition of Endothelial Dysfunction As described above, the vascular endothelium is not merely a passive barrier lining the lumen of blood vessels, it is a dynamic and multifunctional organ that actively regulates vascular tone, coagulation, inflammation, and permeability. The loss of endothelial homeostasis is a common denominator in the onset and progression of numerous diseases, including atherosclerosis, hypertension, diabetes, and sepsis. This pathological state, referred to as endothelial dysfunction, acts as both a trigger and an accelerator of disease processes. Traditionally, endothelial dysfunction has been defined as an imbalance between vasodilatory and vasoconstrictive factors, often characterized by decreased NO production and bioavailability, the presence of oxidative stress, and increased production of cyclooxygenase-2 (COX)-derived prostaglandins, endothelin-1, and/or angiotensin II. These alterations promote a pro-thrombotic environment, increase the permeability and leakage of the endothelial barrier, and heighten inflammatory responses. However, as described above, recent advances in single-cell transcriptomics, metabolomics, chromatin accessibility assays and other technologies have revealed the existence of diverse EC subpopulations with distinct phenotypes and remarkable plasticity across physiological and pathological contexts. Considering these new findings, and based on the consensus guidelines presented in this document, we propose an updated definition. Building upon this foundation, we now recognize endothelial dysfunction as also involving chronic and maladaptive shifts in EC phenotype, driven by environmental and genetic factors that impair EC plasticity, disrupt cellular homeostasis, and contribute to vascular inflammation, remodeling, and disease progression. Therefore, our recommended updated definition of endothelial dysfunction is provided in Table 1 . Table 1: Updated Definition of Endothelial Dysfunction A state characterized by an imbalance between vasodilatory and vasoconstrictive factors (such as decreased vasodilator and/or increased vasoconstrictor factors), along with heightened pro-inflammatory, pro-oxidative and pro-thrombotic responses and/or dysregulation of EC barrier permeability. It may also involve chronic and maladaptive phenotypic switching of ECs, marked by a loss of plasticity and function. These features occur along a continuum and may manifest in combination depending on the disease context. Together, these alterations contribute to persistent vascular damage, remodeling, and the genesis and/or progression of cardiovascular and metabolic diseases. Open in a new tab The involvement of endothelial dysfunction in different pathological states is described below, in brief. These areas represent ongoing efforts to better understand endothelial dysfunction, so targeted therapies can be developed to preserve or restore endothelial function in the hope of preventing or mitigating disease progression. Several in-depth reviews are referenced below that provide more detailed overviews within each area of endothelial dysfunction. Regulation of Vascular Tone and Hypertension There is strong evidence supporting a bidirectional relationship between hypertension and endothelial dysfunction. Accordingly, several studies have demonstrated that endothelial dysfunction can precede the clinical onset of hypertension, supporting the idea that early vascular abnormalities contribute to the initial rise in blood pressure ( 15 ) ( 16 , 17 ). For example, children and young adults with hypertensive parents show reduced endothelial-dependent vasodilation even before developing hypertension, indicating a heritable or early-life predisposition to endothelial dysfunction ( 18 ). Once hypertension is established, elevated blood pressure imposes mechanical stress and circumferential stretch on endothelial cells, which amplifies endothelial dysfunction and structural injury ( 19 ) ( 20 ) ( 21 ) ( 22 ) ( 23 ). This mechanical load acts synergistically with existing endothelial abnormalities, contributing to a vicious cycle in which endothelial dysfunction promotes hypertension, and hypertension further accelerates endothelial damage. Because ECs form the interface between circulating blood and the vessel wall, they are uniquely positioned to detect and respond to changes in blood flow and pressure. They accomplish this through a variety of subcellular structures and protein assemblies that function as mechanosensors ( 24 ). These include ion channels, G protein–coupled receptors (GPCRs), caveolar domains, receptor tyrosine kinases, the glycocalyx, primary cilia, junctional proteins, integrins, and elements of the cytoskeleton. Once these mechanosensitive structures are engaged, ECs activate mechanotransduction pathways that modify cell shape, regulate gene expression, and adjust functional responses. These downstream signals coordinate the release of vasodilatory and vasoconstrictor mediators, enabling communication between ECs and VSMCs. Alterations or dysfunction within this mechanosensing machinery can exacerbate blood pressure elevation, primarily by disrupting the regulation of vascular tone. Below, we provide a brief overview of the key mechanisms that become disrupted during endothelial dysfunction in hypertension and that contribute to abnormal vascular responses. Endothelial dysfunction occurs in many vascular beds during hypertension and is strongly associated with reduced NO synthesis and bioavailability, due to perturbations in the nitric oxide synthase (NOS) expression, activity, and coupling, as well as increased production of ROS, also known as oxidative stress. As a consequence of the NO/ROS imbalance, an impaired vasodilation, enhanced vasoconstriction, and elevated myogenic tone, which can reduce vascular conductance and compliance, increase vascular resistance, and structural remodeling, all of which contribute to sustained high blood pressure. In large arteries, such as the aorta, where NO plays a significant role, disturbances in the NO/ROS balance also lead to increased pulse wave velocity. Importantly, the role of NO, as an endothelium-derived factor diminishes as the lumen diameter decreases in most arteries ( 25 ) (with some exceptions, such as the pulmonary arteries). In contrast, EDH plays a more prominent role in smaller arteries, which will be discussed in detail later ( 25 ). In combination with increased ROS and decreased NO bioavailability, endothelin-1 levels are elevated in hypertension, which further contributes to the enhanced vascular tone and remodeling observed in the disease ( 26 ). Several studies have highlighted the role of endothelial-derived prostanoids in hypertension, particularly noting an imbalance characterized by increased vasoconstrictor prostaglandins (e.g., PGE 2 ) and decreased vasodilatory prostacyclin (PGI 2 ). This shift contributes to elevated vascular tone and blood pressure. Professor Vanhoutte has conducted extensive research on the role of endothelial-derived prostanoids in hypertension ( 27 ). His studies have demonstrated that in hypertensive conditions, there is an upregulation of COX-derived vasoconstrictor prostanoids, such as prostaglandin H 2 (PGH 2 ) and thromboxane A 2 (TXA 2 ), which contribute to endothelium-dependent contractions and increased vascular tone. This imbalance between vasodilatory and vasoconstrictive prostanoids plays a significant role in the pathophysiology of hypertension. However, it is important to note that, in conditions where the endothelium produces high levels of PGI 2 , this prostaglandin could interact with TP receptors (TXA 2 receptors) and induce vasoconstriction instead of vasodilation ( 28 ) More recent evidence suggests that increased sphingolipids, such as ceramide, are produced by ECs and can stimulate ROS production ( 29 ). Studies have shown that hypertensive patients have increased plasma sortilin levels, which can increase ceramide levels ( 30 ). This sortilin-dependent switch in ceramide homeostasis has the potential to switch flow-dependent dilation from being NO- to hydrogen peroxide-dependent ( 31 ). Despite eliciting a flow-dependent dilation, hydrogen peroxide is linked with pro-inflammatory and atherogenic pathways, thereby contributing to the inflammation and remodeling observed in hypertension. Control of Coagulation and Thrombosis The endothelium plays a crucial role in maintaining a balanced hemostatic environment by expressing both pro-thrombotic and anti-thrombotic factors (e.g., vWF, thrombomodulin, tPA). Under healthy conditions, factors that inhibit clot formation predominate, thus preventing unwanted thrombosis, and ensuring that clotting occurs only when needed. In disease states, for example, in atherosclerosis, the dysfunctional endothelium can shift the balance towards a pro-thrombotic state and express adhesion molecules and pro-coagulant factors, facilitating platelet adhesion and aggregation. This contributes to plaque formation and the risk of thrombosis, leading to events such as myocardial infarction or stroke. vWF plays a key role in initiating platelet adhesion and stabilizing factor VIII, but its excessive release by dysfunctional endothelium, possibly driven by increased superoxide formation ( 32 ) and the presence of ultra-large multimers, can tip the balance toward thrombosis ( 33 ). Simultaneously, dysregulation of anti-thrombotic molecules, such as thrombomodulin ( 34 ), tPA ( 35 ), NO, and prostacyclin, shifts the endothelium toward a pro-thrombotic state, contributing to vascular diseases. Barrier Function and Permeability The single layer of ECs lining the lumen of the artery acts as a selective barrier that regulates tissue perfusion to maintain organ homeostasis. EC injury and dysfunction can compromise this barrier function, leading to increased vascular permeability. This “leakiness” is a hallmark of inflammatory diseases, contributing to tissue edema and, in severe cases like acute respiratory distress syndrome or septic shock, to multi-organ dysfunction. ECs are connected by tight junctions (occludins and claudins) and the vascular endothelial cadherin (VE-cadherin). These proteins are crucial for connecting adjacent ECs and maintaining the barrier function. Exposure to molecules such as histamine, VEGF, or thrombin can reversibly disrupt VE-cadherin-mediated junctions in ECs of the microcirculation, leading to an acute increase in vascular permeability. During inflammation, however, Src activation, which can occur through a variety of pathways, including increased ROS, can internalize VE-cadherin, thereby destabilizing the connection between ECs and increasing vascular permeability ( 36 ). Inflammation and Immune Cell Recruitment Under healthy conditions, the endothelium regulates immune surveillance by controlling the passage of leukocytes into tissues through the expression of selectins, integrins, and adhesion molecules (like VCAM-1 and ICAM-1). Inflammatory signals increase ROS production and upregulate VCAM-1, ICAM-1 and MCP-1. This promotes the recruitment, and para- and trans-migration of immune cells into the vessel wall, which under healthy conditions is an essential part of the immune response. Increased adhesion molecule expression leads to more leukocyte recruitment. Endothelial dysfunction can lead to aberrant expression of these proteins, leading to a sustained and chronic inflammatory environment that further damages the endothelium and surrounding tissues. Furthermore, the endothelial glycocalyx has the ability to prevent leukocytes and platelets from adhering to the cell surface. In endothelial dysfunction, the protective glycocalyx is compromised due to enzymatic degradation, inflammatory mediators, and oxidative stress, thereby increasing the ability of leukocytes to adhere to the cell surface and ultimately migrate ( 37 ). The activated leukocytes, in turn, can produce additional ROS and pro-inflammatory cytokines, creating a feedback loop that sustains and amplifies inflammation. This sustained leukocyte infiltration is central to the development of conditions like atherosclerosis, where monocytes adhere to and infiltrate the vessel wall, eventually transforming into foam cells and contributing to plaque formation. Response to Infectious Agents and Systemic Infections Pathogens and damage-associated molecular patterns (DAMPs) are detected by ECs through pattern recognition receptors such as toll-like receptors ( 38 ). As such, ECs act as “sentinel cells” capable of initiating early immune responses by regulating endothelial barrier function and leukocyte migration, as described above. In severe infections (e.g., sepsis or COVID-19), the endothelium can be damaged by both the pathogens and the host’s inflammatory response. This injury can lead to severe endothelial dysfunction, resulting in coagulopathy, increased permeability, and disseminated intravascular coagulation, all of which can culminate in multi-organ failure ( 39 ). Role in Angiogenesis ECs play an essential role in balancing angiogenesis and rarefaction of blood vessels. As such, through the release of VEGF and other pro-angiogenic factors, such as pituitary adenylate cyclase-activating polypeptide (PACAP) and platelet-derived growth factor (PDGF), ECs control the formation of new blood vessels, which is crucial during wound healing and tissue regeneration. VEGF receptors on tip cells dimerize and autophosphorylate when VEGF binds, leading to EC proliferation and migration. In malignant tumors, cancer cells exploit angiogenic pathways to form new blood vessels by expressing hypoxia-inducible factor 1 (HIF-1) and secreting VEGF. ECs subsequently found within the tumor vasculature display several differences compared with healthy ECs. Primarily, these tumor ECs become unresponsive to several inflammatory stimuli, which would normally increase leukocyte migration ( 40 ). For instance, tumor ECs display reduced expression of ICAM-1, ICAM-2, VCAM-1 and E-selectin to attenuate leukocyte binding. They also contain an increased abundance of VEGF receptors (VEGFR1 and 2), to promote EC survival and proliferation ( 41 ). Similarly, aberrant angiogenesis can often be attributed to diminished NO bioavailability, which is required to mediate the pro-angiogenic effects of VEGF, thereby promoting microvascular rarefaction ( 42 ). In diabetic retinopathy, for example, this leads to fragile and leaky vessels in the eye, contributing to vision loss ( 43 ). Metabolic Regulation and Diabetes Obesity, diabetes, and insulin resistance lead to endothelial dysfunction. Hyperglycemia causes damage to the ECs through a variety of mechanisms, including the increased glucose flux through the polyol pathway, promotion of the non-enzymatic glycation of proteins, lipids, and nucleic acids, leading to the formation of Advanced Glycation End-Products (AGEs), augmented protein kinase C (PKC) activation, and over activation of the hexosamine pathway. The production of AGEs represents one of the key mechanisms of endothelial dysfunction in diabetes. AGEs interact with receptors for advanced glycation end-products (RAGE) on ECs and the AGE-RAGE complex activates NFκB signaling that further enhances ROS production and inflammation ( 44 ). Increased peroxynitrite formation is now considered to be strongly linked with endothelial dysfunction in diabetes ( 45 ). Peroxynitrite, formed when superoxide anion interacts with NO, can degrade several protein complexes to produce nitrotyrosine. Diabetic patients are known to have increased plasma nitrotyrosine ( 46 ). As well as acting as a ROS on vascular tone and reducing NO bioavailability, peroxynitrite initiates lipid peroxidation on LDL, leading to the formation of the pro-atherogenic oxLDL. Enhanced oxLDL can augment foam cell formation and plaque deposition in arterial walls. Collectively, these changes contribute to both micro- and macrovascular complications associated with diabetes (e.g., retinopathy, nephropathy, neuropathy) and enhanced cardiovascular disease risk. 2. THE ASSESSMENT OF ENDOTHELIAL FUNCTION Preparation and Composition of Experimental Solutions for Isolated Vessels Experiments ( ex vivo ) Rather than perfusing tissue with blood during ex-vivo investigations, most experimental studies use simplified physiological saline solutions. These solutions are much easier to prepare, store, and use, and their variation over time can be kept to a minimum. However, the composition of physiological saline solutions must be carefully considered in order to best mimic the investigated in vivo physiological or pathophysiological conditions. A standard recipe for a widely used physiological saline solution is presented in Table 2 . When studying arteries from most mammals, this solution should be heated to 37°C, bubbled vigorously with 5% CO 2 /balance air, and adjusted to pH 7.40 before use. Table 2. Composition of a physiological saline solution Concentration (mM) Na + 141 K + 4.0 Mg 2+ 1.2 Ca 2+ 1.6 Cl − 121.8 HCO 3 − 22 SO 4 2− 1.2 H 2 PO 4 − /HPO 4 2− 1.18 Glucose 5.5 EDTA 0.03 Open in a new tab Typical composition of a physiological saline solution suited for ex vivo evaluation of endothelial function. The solution should be heated, aerated with a gas mixture containing 5% CO 2 /balance air (or other appropriate O 2 level), and adjusted to 7.40 pH. Note that although phosphate in the displayed recipe is added as H2PO 4 − , after equilibration and titration to pH 7.40, more than half will be present as HPO 4 2− because the pKa value is 7.21. EDTA, ethylenediaminetetraacetic acid. Small deviations will occur when pH is titrated using HCl or NaOH. When blood vessels are mounted for isometric investigation in wire myographs, their luminal and abluminal surfaces are in contact with the same solution. In pressure myographs, it is possible, although not common, to use solutions of different composition in the lumen and bath compartment, for instance, by perfusing only the lumen with an albumin-containing solution. Please refer to the Guidelines for Measurement of Vascular Function and Structure in Isolated Arteries and Veins ( 1 ) for detailed instructions on preparing solutions for pressure myographs. We recommend the indicated composition ( Table 2 ) of the experimental solution for isolated wire and pin myographs. The saline solution described in Table 2 matches the basic ionic composition of plasma and interstitial solution. However, as discussed in detail below, investigators should be aware of important shortcomings that result from the reductionist approach of using simple saline solutions. These solutions lack the formed elements of blood (i.e., erythrocytes, leukocytes, and thrombocytes) and are without important molecules from plasma and interstitial solution (e.g., albumin, fatty acids, amino acids, and lactate), which may influence cellular functions and the experimental results. Further, although Table 2 presents the solution that best approximates physiological conditions and represents the most broadly accepted standard, readers will encounter several additional solutions referenced throughout the manuscript. These formulations are not alternatives to the standard PSS, but are technique-driven adaptations used in particular experimental contexts, such as electrophysiology, freshly isolated EC preparations, lymphatic studies, and others, where the unique requirements of the method necessitate deviations from the physiological range. O 2 Choosing an appropriate experimental O 2 concentration for ex vivo studies is not straightforward because saline solutions lack the O 2 -carrying capacity offered by hemoglobin in blood. Systemic arterial blood from healthy human individuals has a PO 2 around 100 mmHg, whereas interstitial solution in various organs differs in PO 2 from around 8 mmHg in the superficial skin to 29 mmHg in skeletal muscle, 34 mmHg in the brain, 58 mmHg in the intestine, and 72 mmHg in the kidney ( 47 ). At rest, human mixed venous blood has a PO 2 of approximately 40 mmHg; however, the partial pressure will drop during exercise and in pathophysiological conditions where O 2 extraction is increased. For simple preparations of isolated arteries, a gas mixture containing 5% CO 2 /balance air is usually employed. This gas mixture contains around 20% O 2 corresponding to a PO 2 of approximately 143 mmHg in a humidified atmosphere. There are also examples of investigators using 5% CO 2 /balance O 2 , which corresponds to a PO 2 of around 677 mmHg, however, such supraphysiological O 2 concentrations can harm the tissue ( 48 - 50 ). It may be particularly detrimental when elevated O 2 levels are combined with experimental procedures, such as microscopy utilizing laser excitation, that can increase the generation of ROS. Although solutions equilibrated with 95% O 2 should generally not be used for isolated arteries or cells, they can be meaningful for more complex preparations, like isolated perfused hearts or blood vessels covered by substantial perivascular tissue, in order to compensate for the longer diffusion distances and avoid development of tissue hypoxia due to the low O 2 -carrying capacity in absence of hemoglobin. For these more complex cardiovascular preparations, one must weigh the risk of regional hyperoxia against the risk that deoxygenation of the saline solutions can cause tissue hypoxia in other regions. Recent studies have employed artificial O 2 carriers to improve oxygenation of isolated perfused hearts, and such particles could represent a better way to replicate the O 2 concentrations and partial pressures observed in vivo ( 51 ). Some vascular preparations require particular attention to O 2 levels: Systemic veins, lymphatic vessels, and pulmonary arteries typically experience lower luminal O 2 levels in vivo compared to systemic arteries. CO 2 /HCO 3 − and alternative buffers The CO 2 /HCO 3 − buffer system in most physiological solutions is not only the most important for minimizing immediate changes in pH, it also acts as a mobile buffer that accelerates transfer of H + equivalents between sites of production, consumption, and transport ( 52 ). More recently, it has been appreciated that HCO 3 − is sensed by ECs of resistance arteries both in intracellular compartments and at the plasma membrane ( 53 ) ( 54 , 55 ). It is therefore important that CO 2 and HCO 3 − are maintained at levels appropriate for the evaluated physiological or pathophysiological condition. Systemic arterial blood typically contains 40 mmHg CO 2 whereas mixed venous blood has a PCO 2 of around 46 mmHg. Traditionally, physiological saline solutions are aerated with a gas mixture containing 5% CO 2 corresponding to a PCO 2 of 36 mmHg in a humidified atmosphere. To achieve a pH of 7.40 at 37°C, this concentration of CO 2 balances a HCO 3 − concentration of 22 mM ( Table 2 ). If a more physiological CO 2 level of 40 mmHg is used, the concentration of HCO 3 − should be elevated to 24 mM. Alternative HCO 3 − concentrations are relevant for some organisms, e.g., fish ( 56 ), and also if experiments are performed at lower temperatures than 37°C because both the solubility of CO 2 , the pKa of the CO 2 /HCO 3 − equilibrium, and the saturated water vapor pressure are temperature dependent ( 57 ). It is important to remember that pH of the experimental solutions should be adjusted after addition of all buffer components, when the solutions have been aerated long enough to reach CO 2 /HCO 3 − /H + equilibration, and after heating to 37°C. Under some experimental conditions, e.g., during microelectrode-based recordings where mechanical disturbances can influence the recordings, it can be difficult to continuously bubble solutions with the vigor required to achieve a concentration of 5% CO 2 . Under these conditions, including HEPES or other artificial buffers, can improve pH control. Artificial buffers are also required if ion substitution experiments (e.g., replacing HCO 3 − with Cl − and omitting CO 2 ) are performed to explore the functional importance of the CO 2 /HCO 3 − buffer system ( 58 ). It should be noted, however, that these alternative buffers can have their own vascular effects ( 59 ). K + The activity of excitable tissue influences the concentration of K + in the interstitial solutions surrounding intra-organ blood vessels. The K + concentration is particularly dynamic in the interstitium of the heart and skeletal muscle where it easily reaches 10 mM during exercise and ischemia ( 60 , 61 ). Still, for most studies aiming at resembling basal conditions, it is desirable to use a concentration of K + that mimics that of arterial plasma at rest, which is around 4 mM. Ca 2+ Many previous studies of isolated vascular tissue have used supraphysiological concentrations of Ca 2+ , often around 2.1-2.6 mM, which is the range of the total Ca 2+ concentration (i.e., ionized and bound) in plasma. Elevated free Ca 2+ concentrations have most likely been chosen in order to enhance tension development and thus improve the signal-to-noise ratio, but they could influence vascular function inappropriately. Choosing a proper Ca 2+ concentration, however, is not trivial as even in protein-free saline solutions, the Ca 2+ activity is less than the total Ca 2+ concentration. Previous measurements from Krebs-Ringer HCO 3 − buffers show that the Ca 2+ activity is around 20% lower than the total Ca 2+ concentration in protein-free buffers ( 62 ). Therefore, we suggest adding 1.6 mM Ca 2+ to reach an ionized Ca 2+ level in the physiological range of 1.2-1.4 mM. Energy sources: glucose, fatty acids, amino acids, and lactate Glucose is often the only nutrient included in physiological saline solutions ( Table 2 ). Because fatty acids are the main energy source in many resting cell types, and amino acids and lactate are also oxidized for energy production under some circumstances, omission of these important nutrients is potentially problematic. Investigators have previously compensated for the absence of other nutrients by elevating the glucose concentration above physiological levels. Whereas additional glucose might be helpful for avoiding cell starvation, it should be done with caution since elevated glucose concentrations, as observed during diabetes mellitus, may contribute to the development of vascular dysfunction ( 63 ). Adding longer-chain fatty acids to simple saline solutions is complicated by their low aqueous solubility, but the solubility is improved if albumin-containing solutions are employed ( 64 ). Although absence of amino acids in physiological saline solutions may not be a problem for shorter-lasting experiments, the addition of amino acids should be considered for experiments of longer duration. If ECs or blood vessels are maintained in culture, the use of complete culture media is essential to provide the nutrients and chemical building blocks required for maintained endothelial function. Amino acids are essential for key EC processes, including NO synthesis from arginine, and addition of arginine has been found to improve vasorelaxation ex vivo ( 65 ). Viscosity Due to the absence of formed elements and protein (e.g., albumin and fibrinogen), simple physiological saline solutions, like the one described in Table 2 , have low viscosity compared to blood. Adding albumin to increase the viscosity of the solutions is relevant when studying, for instance, flow-mediated vasorelaxation where shear stress is tested across the physiological range ( 66 ). Although the addition of albumin, as described above, can also help carry fatty acids, it carries experimental challenges. Aeration of albumin-containing saline solutions generates foam that can disturb experimental recordings. It should also be considered that the free concentration of added drugs can be lowered in albumin-containing solutions due to protein binding. Antioxidants The oxidative state influences many processes and drugs relevant to cardiovascular function. Addition of antioxidants to physiological solutions, for instance, modifies the reuptake of noradrenaline and hence the elicited contractile response ( 67 ). In the solution suggested in Table 2 , ethylenediaminetetraacetic acid (EDTA) is added at a low concentration that does not contribute substantially to Ca 2+ -buffering but provides antioxidant effect ( 68 ). Adding more relevant physiological antioxidants could be considered. The solution described here is the most common and widely used, closely matching physiological conditions; however, some groups use alternative buffers, such as 3-(N-morpholino)propanesulfonic acid (MOPS)-based solutions, which maintain stable pH without the need for continuous bubbling, simplifying preparation and experimental setup for arterial studies but potentially altering intrinsic contractile activity in lymphatics. As noted by the authors of these guidelines, it is important to consider these different solutions when designing experiments to ensure reproducibility and comparability of results across studies. Most Used Techniques for Assessing Endothelial Function Please refer to the Guidelines for Measurement of Vascular Function and Structure in Isolated Arteries and Veins ( 1 ) for detailed instructions on mounting large and small vessels using pin, wire, or pressure myograph systems. This reference can also be used to guide the evaluation of VSMC contractility. Investigators should be aware that different euthanasia approaches can influence vascular reactivity. For example, CO 2 exposure and certain anesthetic agents have been reported to alter endothelial function, modify nitric oxide bioavailability, or affect smooth-muscle responsiveness. Therefore, laboratories should use a method approved by their institutional guidelines and, most importantly, maintain consistency within studies to avoid introducing variability attributable to the euthanasia procedure. The sections below will focus exclusively on the assessment of endothelial function, assuming that muscle viability has been previously confirmed. A. Pin Myograph for Large Arteries The classical paper by Furchgott and Zawadzki ( 2 ) first reported the obligatory role of ECs in acetylcholine (Ach)-induced relaxation of large arteries such as rabbit aorta, superior mesenteric and pulmonary arteries. Since then, isolated arteries mounted in a myograph setup have been widely used to assess the endothelial vasorelaxation function. Arterial tubular rings and strips can be mounted in a glass-chamber tissue organ bath or in a wire myograph. Fine stainless steel or tungsten pins, L-shaped bars, or hooks are passed through the lumen of the blood vessel. Importantly, while tungsten (wire or pins) is widely used, researchers should be aware that oxidation of tungsten has been reported to alter endothelial function ( 69 ). Subsequently, the vessel is stretched to a level that approximates physiological conditions. This resting tension is specific for each vessel type, animal model (e.g., human, rodent models), and myograph system used, allowing the vessel to produce maximal active isometric force in response to a pharmacological stimulus ( 1 ). This method has been used to evaluate endothelium-dependent responses in isolated mammalian arteries and veins (including human and murine), as well as in other vertebrates such as reptiles, amphibians, and fish. The pin myograph setup does not allow for the evaluation of flow/shear stress-induced endothelium-dependent relaxation. Therefore, pharmacological agents that increase intracellular Ca 2+ concentration [Ca 2+ ] i in ECs have been used as pharmacological tools to assess endothelium-dependent relaxation in isolated arteries via myography. These include muscarinic agonists (e.g. ACh, methacholine), bradykinin and the Ca 2+ ionophore A23187. A healthy endothelium is important for the vasorelaxation to several hormones and local factors such as angiotensin 1-7, adiponectin, VEGF, PGI 2 , glucagon-like peptide-1 (GLP1), insulin, oxytocin, estrogen, among others ( 70 ). As described earlier, in larger arteries, such as the aorta and superior mesenteric artery, NO has been identified as the major endothelium-derived relaxing factor ( 25 )( 71 , 72 ). The concentration response curve to ACh is the most commonly used protocol for evaluating endothelium-dependent relaxation in myograph/organ bath setup. When applied to healthy vessels using wire/pin myography in the absence of pre-constricting agent, ACh does not elicit a significant relaxation response ( 73 ). Therefore, a sustained contraction must be induced before assessing the endothelium-dependent relaxation curve. Alpha (α)1-adrenergic receptor agonists (such as phenylephrine and methoxamine) have been effectively used in large vessels for this purpose given the widespread distribution of α 1 -adrenergic receptors in vessels and to avoid limited contraction mediated by α 2 - and/or β-adrenoceptors. U46619, a TXA 2 receptor (TP receptor) agonist, is known to induce a potent and stable contraction, which can be advantageous for vessels with low or unstable vasoconstriction activity. However, TP receptor signaling may promote ROS production ( 74 ). Consequently, U46619-induced contraction may lead to a decrease in NO bioavailability, potentially resulting in a reduced endothelium-dependent relaxation response. Additionally, U46619 could block the contractile response elicited by ACh through the release of TXA 2 in some cardiovascular diseases. Thus, in certain circumstances, serotonin and endothelin-1 are also agonists that have been used. The choice of the agonist for inducing precontraction needs to consider vessel type, animal species, sex, and the experimental model (e.g. a model of cardiovascular disease). The precontraction needs to be sufficient to allow a measurable relaxation response, but if it is too strong, the vessel may not relax properly. For large arteries, a target of approximately 50-80% of the maximum high KCl-induced vasoconstriction is typically used to achieve submaximal contraction ( 71 ). However, this range can vary depending on several factors, including the presence of perivascular adipose tissue, sex, age, disease, and vessel length. Therefore, we recommend that the individual performing the experiment has experience in artery mounting to ensure that the control group (healthy arteries) consistently achieves precontraction within the 50-80% range, as shown in Fig. 4 . Importantly, in experimental groups, this range may shift even when using similar concentrations of common vasoconstrictors (e.g., phenylephrine, U46619, etc.). For example, arteries from diseased animals may exhibit altered agonist sensitivity compared to healthy arteries. Conversely, arteries from older animals or those with severe hypertension may show reduced contractility due to increased collagen deposition (i.e., stiffness) or a phenotypic switch of smooth muscle cells to a less contractile state. Given these variations, we recommend generating a concentration response curve to the selected agonist for each novel experimental group. This will help determine the appropriate submaximal contraction range (50-80%) specific to that group, which may differ from the control. Vasodilator responses are usually calculated as percentage of relaxation relative to the agonist-induced precontraction [(precontraction- response to ACh)/(precontraction - resting tone)]. Alternatively, it can be expressed as a percentage of the maximal relaxation elicited by papaverine. Figure 4. Open in a new tab Force recording of mouse aorta. A maximal contraction was induced by applying 80 mM KCl. Following wash, U46619 was applied to induce submaximal contraction (70%). A concentration response curve to acetylcholine (ACh) was obtained. Figure 4 exemplifies a protocol for evaluating endothelium-dependent relaxation in mouse aorta. Accordingly, an aortic ring (2 mm) from a male mouse was mounted in a pin myograph under a basal tone of 5 mN. Eighty mM KCl was applied to induce maximal contraction. Of note, when using high-concentration KCl in physiological solution to induce vascular contraction, it is recommended to replace an equivalent amount of NaCl with KCl so that the final solution remains isoosmotic, thereby eliminating potential confounding effects from altered osmolarity. However, it is important to emphasize that replacing NaCl with KCl alters the transmembrane Na + gradient and therefore affects both Na + -Ca 2+ exchange (NCX) and/or Na + -H + exchange (NHE), which should be acknowledged when interpreting vascular responses under these conditions ( 75 ). After several washes, the vessel was precontracted with U46619 until reaching ~60% of the KCl-induced vasoconstriction. Then, a concentration-response relaxation curve to ACh was performed. Concentration-response curves become sigmoidal when plotted on a semilog scale (log concentration vs . response), making it the standard method for determining potency (EC 50 ) and/or maximum response (E max ) [for further information please see reference ( 1 )]. ACh-induced relaxation was calculated as percentage of relaxation relative to the agonist-induced precontraction, generating the ACh-induced relaxation curve. ACh (1 μM) produces 80-100% relaxation in submaximally precontracted rat aorta, correlating with 60-75% of the ECs being retained ( 2 ). In large mouse arteries, a relaxation response to ACh (10 μM) above 60% indicates endothelial integrity ( 71 , 76 , 77 ). Mechanical removal of the endothelium in large vessels, such as aorta can be performed by gently rubbing the lumen with a needle, a cocktail stick, a jeweler forceps, or by rotating the arterial ring around the pin wire in a myograph ( 78 )( 79 ). Chemical denudation using detergents such as 3-[(3-cholamidopropyl)-dimethyl-ammonio]-1-propanesulfonate (CHAPS) is another option ( 80 , 81 ) though it carries a higher risk of damaging VSMCs. When endothelium removal is successful, ACh fails to induce relaxation. It is important to note that, in the aorta, basal NO appears to be more susceptible to destruction by superoxide anion than NO produced in response to ACh ( 82 ). Therefore, to investigate the role of basal endothelial-derived factors on vascular tone, the anticontractile effect of the endothelium can be assessed. A concentration-response curve to contractile agonists like phenylephrine can be obtained in arterial rings with and without endothelium ( 83 ). B. Wire Myography for Small Arteries Wire myography can be used to determine the effect of EC signaling on the vascular tension of small arteries. Any endothelium-dependent effect can be confirmed by endothelium denudation, performed as described in Wenceslau et al. ( 1 ). Similar to the pin myograph for large arteries, to study an endothelium-dependent effect in the wire myograph, the presence of a functional endothelium must first be confirmed. This is often performed on precontracted artery segments as described above. However, for small arteries, a target of approximately 70-90% of the maximum high KCl-induced contraction is typically used to achieve submaximal contraction. This is because healthy small arteries tend not to maintain vasoconstriction for extended periods when the precontractile response is lower than 70% of the maximum KCl-induced contraction. For healthy small arteries, agonists such as ACh (1-10 μM)/carbachol (1-10 μM) or bradykinin (1 μM) elicit endothelium-dependent vasodilation and should induce >70% relaxation after preconstriction for the arterial segment to be deemed to have an intact endothelium. In the case of denudation, the relaxation to these agonists should be <15%. Please note that these are best-practice recommendations rather than experimentally validated cutoffs, and they should be interpreted within the context of each laboratory’s methodology and experimental conditions. Please see practical tips for endothelium denudation in resistance arteries in Table 3 . Table 3. Mechanical Denudation of the Endothelium in Resistance arteries (most common method): ✓ Dissection & Mounting: Carefully dissect the artery under a stereomicroscope. Avoid stretching or damaging the endothelium during isolation. Mount the vessel onto the wire myograph with stainless steel or tungsten wires (usually 25-100 μm diameter). Importantly, while tungsten (wire or pins) is widely used, researchers should be aware that oxidation of tungsten has been reported to alter endothelial function ( 69 ). Some researchers have used gold-plated tungsten as an alternative ( 69 ) ✓ Mechanical Denudation (most common method): Gently rub the luminal surface: After mounting, a human hair or a fine nylon monofilament (e.g., from a fishing line or surgical suture, wetted in Krebs solution) is threaded through the lumen and gently moved back and forth once or twice to mechanically remove endothelial cells. The hair or filament should be clean and wetted in Krebs buffer to avoid drying the vessel. ✓ Confirmation of Denudation: Assess VSMC contractile capacity using high KCl (typically 80-120 mM). A reduced contraction to KCl compared to healthy, non-denuded vessels indicates possible VSMC damage during the denudation process. After a washout and equilibration period, test with acetylcholine (ACh, 1 μM) following a phenylephrine or U46619 precontraction. A lack of relaxation to ACh (typically <15%) confirms successful denudation. Open in a new tab To investigate the contribution of NO at a basal level and following stimulation (e.g. ACh), NG-Nitroarginine Methyl Ester (L-NAME) or NG-monomethyl-L-arginine (L-NMMA) can be applied to the bath at ~100 μM to inhibit NOS. It is important to note here that L-NAME can inhibit neurogenic dilatation elicited by electrical field stimulation (EFS), whereas L-NMMA has no effect on the EFS-induced neurogenic dilatation ( 84 ). However, L-NMMA has been reported to enhance NO production ( 85 ) and inhibit the basal but not the agonist-stimulated NO-dependent dilatation in the rat aorta ( 86 ). Thus, it is good practice to ascertain whether there are differences in the effect of these NOS inhibitors on the arterial preparation chosen. The effect of basal NO release can be ascertained by performing a concentration-effect curve with a vasoconstrictor such as noradrenaline, phenylephrine or U46619. The level of enhanced contraction, often shown as the shift in EC 50 for resistance arteries, of these vasoconstrictor agonists in the presence of L-NAME or L-NMMA, will give an indication of the basal level of NO release. To determine the level of stimulated NO-induced relaxation, the arterial segment should be preconstricted to 70-90% of its maximal tone with the vasoconstrictor of choice, as described above. A concentration-response curve is then performed with the vasodilator as the control experiment. In parallel, the same experiment is performed on another segment of the same artery, with L-NAME or L-NMMA (100 μM) applied 10-30 mins prior to preconstriction. Care should be taken here to ensure the level of constriction is not significantly enhanced compared to the control segment, and, if necessary, the concentration of vasoconstrictor used for pre-constricting the vessel should be reduced. The shift in EC 50 and/or E max of the vasodilator in the absence compared to the presence of L-NAME or L-NMMA will indicate the NO-mediated component of the relaxation. Of note, we previously highlighted the use of two segments from the same artery, one serving as baseline and the other receiving a pharmacological blocker, as an important element of experimental rigor. However, it is equally important to acknowledge that alternative designs are widely used in the vascular reactivity literature. For example, some studies employ washout periods between consecutive protocols, or a paired-segment approach in which both arterial rings first generate a control response to the agonist. Only one segment is then exposed to the inhibitor, while the second segment functions as a time control. In this design, the control vessel should reproduce the initial dilatory response, verifying that any attenuation observed in the inhibitor-treated segment reflects successful pharmacological blockade rather than time-dependent drift In addition to inhibiting NOS, 1H-[1,2, 4]oxadiazolo[4,3-a]quinoxalin-1-one (ODQ) can be used to inhibit sGC, the primary downstream target of NO in VSMCs. Similar to the L-NAME experiments described above, ODQ is applied to the bath at 0 μM for 10-30 mins prior to preconstriction and the concentration-effect curve of the chosen agonist. If there is no relaxation induced by ACh/carbachol or bradykinin in the control arterial preparations that are known to have healthy endothelium, one must consider whether the endothelium was damaged in the mounting procedure. However, if there is a lack or impraired response to these agonists when applied to arteries from a diseased animal model or certain patient group (and the researcher is experienced with mounting arterial segments with an intact endothelium), it is then possible to determine whether the endothelium in these arterial segments is compromised or the VSMC lack the capacity to relax to NO-stimulation by studying endothelium-independent NO-dependent relaxation. To do this, the arterial segment is preconstricted and increasing concentrations of an NO donor are applied. Four commonly used NO donors are: sodium nitroprusside (SNP; 1 nM-30 μM), S-nitroso-N-acetylpenicillamine (SNAP; 0.1-10 μM), 3-morpholino-sydnonimine (SIN-1; 0.1-10 μM) and and Diethylamine NONOate (DEA; 1 nM-30 μM). When these are applied to preconstricted arterial segments, they will induce relaxation if the VSMCs have maintained their ability to relax ( 87 ). An alternative to using NO donors would be the soluble guanylyl cyclase activator, 1 μM Bay 41-2272, which can induce relaxation of a preconstricted arterial segment, if the sGC-cGMP pathway in VSMCs is not compromised. The combination of assessing the endothelial NO-dependent relaxation and the ability of the VSMC to relax in response to NO allows the researcher to elucidate the underlying cause of an attenuated NO-dependent relaxation. As well as assessing NO-dependent relaxation, it is important to consider the effect of PGI 2 release, since ACh and bradykinin can also stimulate their production from arachidonic acid ( 27 ). These experiments are commonly performed using the COX inhibitor indomethacin (3 μM), applied to the bath prior to preconstriction and the concentration-response curve to the selected agonist. Inhibition of concentration-dependent relaxation by indomethacin reveals the contribution of prostacyclin and certain prostaglandins. Since COX 1 and 2 are both inhibited by indomethacin, it can often be beneficial to use COX-selective inhibitors ( 88 ), for example SC-560 (COX1-selective, 0.1-1 μM) and NS-398 (COX2-selective, 100 μM). As mentioned earlier, the importance of the hyperpolarizing mechanism increases as the vessel size decreases in endothelium-dependent relaxations ( 72 )( 72 ). Therefore, it is important to understand the role of EDH when studying resistance arteries. For this, the role of K + in EDH is studied using a combination of intermediate-conductance Ca 2+ -activated potassium channel (IK Ca ) and small-conductance Ca 2+ -activated potassium channels (SK Ca ) channel blockers. 1-[(2-chlorophenyl)diphenylmethyl]-1H-pyrazole (TRAM-34; 1 μM) is used to block IK Ca and apamin (50 nM) or UCL 1684 (1 μM) is used to block SK Ca , with these compounds often applied together approximately 5-20 min before preconstriction of the arterial segment and application of the vasodilatory agonist. A shift in the concentration-effect curve (EC 50 and/or E max ) to the agonist in the presence of TRAM-34 and apamin/UCL 1684 provides the EDH component attributed to K + release by the ECs. Alternatively, NS309 (6,7-dichloro-1H-indole-2,3-dione 3-oxime), a positive modulator of SK Ca and IK Ca channels (0.1-100 μM) can be applied to a preconstricted artery segment to determine the influence of these EC channels on vasoreactivity ( 89 ). As mentioned previously, H 2 O 2 , under certain conditions, can be produced and cause a relaxation of arteries. In order to determine the H 2 O 2 -mediated dilatation following stimulation (e.g. by ACh or bradykinin), PEG-catalase (300 U/mL) is added to remove intracellular H 2 O 2 ( 90 ). The mechanisms of action of pharmacological agents, interpretation of pharmacological responses, and considerations of sex differences will be discussed later in these guidelines. C. Pressure Myography for Small Arteries Flow-Induced Dilation (FID): The ability of an artery to sense an increase in blood flow and respond through the production of endothelial-derived mediators is critical to maintain proper tissue perfusion. Determining whether the force of shear stress not only elicits dilation but also generates vasoactive substances that promote a robust vascular environment, may be considered the most physiological approach to assessing the health of the endothelium. Unlike endothelial dysfunction in large conduit arteries which manifest as reduced flow-mediated dilation (FMD) due to a lower bioavailability of NO, the microvasculature can compensate for loss of NO-mediated dilation by utilizing other endothelial-derived mediators such as H 2 O 2 ( 6 ). For clarification, the term flow-induced dilation (FID) will be used when discussing the assessment of flow in isolated arteries as FMD is commonly used to describe the response of a large human conduit artery (e.g. brachial artery) to increased flow in vivo immediately following a period of distal limb occlusion ( 91 ). Essentially, the entire vasculature, from conduit vessels to resistance arterioles, responds to increased shear along the intimal wall by releasing endothelial-derived mediators to elicit dilation, albeit the occasional exception ( 92 ). To date, the majority of studies examining mechanisms of FID have been performed in isolated resistance arteries by leveraging the pressure myography technique. Preclinical studies typically utilize mesenteric arteries from rodents ( 93 ), however other animal models have greatly contributed to our understanding of microvascular FID ( 94 ). Measuring vasoreactivity to flow in human arteries can be achieved by collecting discarded surgical specimens such as adipose ( 95 ) or atrial tissue ( 96 ) as well as performing gluteal biopsies in human participants ( 97 ). Once the tissue of interest is obtained, resistance arteries with internal lumen diameters of ~80-200 μm are identified and dissected. A pressure myography system where both ends of the artery are cannulated onto glass micropipettes of matched resistance is then used for the vascular reactivity experiment. For more details regarding the dissection and cannulation process as well as logistics for performing pressure myography for FID, we direct the reader to this reference ( 1 ). Important considerations for conducting, analyzing, and interpreting FID in isolated arteries are discussed below. Establishing Baseline Tone: To assess the ability of an artery to dilate, it is important that baseline tone be established to determine total vasodilatory capacity. The diameter of the vessel at the initiation of equilibration should be recorded as well as the final diameter post-equilibration. This will inform the investigator whether spontaneous tone has developed. Microvessels from animals typically develop spontaneous myogenic tone which is observed as a reduction in vessel diameter during equilibration of typically 30-50%. This is in sharp contrast to arteries dissected from human specimens (e.g. adipose and atrial tissue) which usually require the vessel to be pre-constricted with a pharmacological agent to reduce the diameter by the same percentage. This difference is likely due to a combination of factors, including prolonged ischemia or extended handling time between tissue excision and the start of the experiment, which can impair vascular function; variability in donor health conditions, and differences in tissue storage conditions, including cold storage and transport buffers, which may reduce myogenic responsiveness. Additionally, intrinsic species differences may also contribute to this phenomenon. The vasoconstrictive endothelial-produced peptide endothelin-1 (ET-1) is commonly used for precontract human species in this situation. For this, it is recommended to start at concentrations of 0.2 nM. However, prior to administration of ET-1, the investigator should calculate the expected diameters following exposure to the vasoconstrictor, which will assist in avoiding over-constriction and possibly damage. For instance, if the lumen diameter of the vessel post-equilibration is 100 μm and did not develop spontaneous tone, (e.g. remains 100 μm in diameter at the end of equilibration) the investigator should calculate and record the desired diameter following constriction with ET-1, which for this example would be 50-70 μm (30-50% of the baseline diameter). Aside from ET-1, other vasoconstrictors have been used for this purpose including but not limited to, norepinephrine (10 μM) and U-46619 (100 nM). If 30-50% constriction is not achieved following the initial concentration, additional concentrations may be necessary to reduce the diameter prior to starting flow. In the event the vessel is constricted beyond 50% of the passive diameter (e.g. 70-80%) the experiment can continue if the lumen of the vessel can still be visualized and perfusate is able to flow through the vessel and elicit a shear response. In the unfortunate circumstance that the vessel over-constricts and a lumen is not visualized, the investigator should consider replacing the organ chamber buffer x3 and allow for relaxation. If this is not accomplished with washing alone, one may consider adding a direct smooth muscle dilator such as papaverine (100 μM), wash, and repeat the pre-constriction step. Finally, if repeated administration elicits only a minimal constriction (<30%) or there is an absence of constriction all together, the investigator should test the viability of the vessel by administering an agent that directly depolarizes VSMC, such as KCl, 60-80 mM, which depolarizes VSMC and produces receptor-independent contraction. A robust contraction to KCl confirms that the VSMC layer is viable, whereas a poor or absent response indicates loss of smooth muscle function and that the vessel is no longer suitable for FID assessment. Evaluating a Control Curve: Once the artery is cannulated, warmed, equilibrated and pre-constricted, vascular reactivity to flow can be measured. This is accomplished by raising and lowering the heights of the open reservoirs in equal and opposite directions, allowing for unidirectional flow through the vessel without increasing intraluminal pressure. However, this may depend on the equipment used; for example, if the investigator has a flowmeter, it can be used to regulate and monitor the flow rate more precisely. Because shear stress depends on both flow rate and vessel diameter, it can be calculated in pressure-myograph preparations when diameter is measured and flow is known or derived from the pressure gradient, ensuring accurate interpretation of the endothelial shear response. The optimal intraluminal pressure in which to perform FID is highly dependent on the species. For instance, human and pig arteries respond best to flow at 60 mmHg ( 94 ), opposed to piglet cerebral vessels (20 mmHg) ( 98 ), rat mesenteric arteries (80 mmHg) ( 99 ), and rabbit pial resistance arteries (30 mmHg) ( 100 ). It is critical that the pipet end diameters are equal (~40 μm) to ensure matched resistance and to avoid fluctuations in intraluminal pressure which will confound the response to flow. Measurements of the internal diameter of the vessel are taken at both 1 min and 5 min (steady state) following a change in the pressure gradient at 5, 10, 20, 50, and 100 cmH 2 O, with most of the dilation (80%) occurring at the 20 cmH 2 O pressure gradient. One needs to be prudent in creating the pressure gradient and changing the heights of the reservoirs as doing so in a rapid manner may change vessel pressure or in some instances result in pressure-induced constriction. It is also important to design the pressure myography system in a way that the perfusing solution is maintained at 37°C regardless of the flow rate and using CO 2 -tight tubing (e.g., Tygon) to make sure PCO 2 and pH will not change and influence tone when flow is manipulated. Importantly, net tone can also be explicitly determined by measuring the passive diameter of the vessel in Ca 2+ -free PSS and expressing active diameter changes relative to this passive state. Cannulated arteries can typically be used for a maximum of two complete experimental protocols on the same vessel; however, reproducibility assays should be performed initially to demonstrate the long-term viability of the preparation. The first experiment serves as a control to assess the overall ability of the resistance arteriole to dilate in response to flow, which, on average results in dilation to ~80% of maximal diameter. The equation to calculate % maximal dilation is provided below. The starting diameter refers to the internal diameter of the vessel prior to initiating flow and after pre-constriction, whereas the maximal diameter is the largest diameter of the vessel at any point during the experiment, typically observed following administration of papaverine. Inner diameter is the preferred and recommended measure for FID, as it directly reflects changes in the lumen in response to flow. In contrast, outer diameter does not directly reflect lumen changes and can vary due to wall thickness, making inner diameter a more functionally relevant parameter for FID. However, some groups use the outer diameter for mesenteric arteries, particularly when additional structural analyses are planned. If outer diameter is used, it must be applied consistently throughout the experiment and analysis to ensure comparability. % Maximal Dilation = [ Diameter at Pressure Gradient − Starting Diameter ¯ ] [ Maximum Diameter − Starting Diameter ] In the event the vessel was exposed to an agent or insult, an overall reduction in vasodilatory magnitude may be observed suggesting vascular dysfunction. Following measurement at the maximal pressure gradient (100 cmH 2 O), the reservoirs are returned to equal heights and allowed to rest at 60 mmHg prior to the next experiment. Inhibitors can be added to the organ bath to determine the mechanism of dilation. There are numerous pharmacological agents that can be utilized to decipher which endothelial-derived compound(s) contributes to FID. Similar to pin and wire myographs, common inhibitors used include but are not limited to: 1) L-NAME, 100μM and 2-(4-carboxyphenyl)-4,5-dihydro-4,4,5,5-tetramethyl-1H-imidazolyl-1-oxy-3-oxide, monopotassium salt (c-PTIO, 1 μM) to assess the contribution of NO; 2) PEG-catalase, 500 U/mL to determine the contribution of H 2 O 2 ; 3) indomethacin (3 μM) to assess the role of COX-derived prostaglandins; and 4) TRAM-34 and apamin/UCL 1684 for EDH. If exposure to one inhibitor reduces FID but residual dilation remains, additional agents may be added to determine the vasoactive mediator responsible for the remaining vasodilatory capacity. In the event that two inhibitors result in complete loss of dilation in separate experiments, it is possible that one vasoactive mediator relies on production of the other to cause vessel relaxation ( 101 ). On the flipside, if multiple inhibitors fail to reduce FID separately (e.g. vessel dilates in the presence of both L-NAME and PEG-catalase) but when combined they reduce the vessel’s ability to dilate (e.g. L-NAME + PEG-catalase) it may signify that one vasoactive compound compensates for loss of the other ( 102 ). Please see illustration in Fig. 5 . Aside from the various inhibitors that can be added to the organ bath to identify endothelial-generated FID mediators produced during flow, an endless list of pharmacological agents can be leveraged to provide mechanistic insight into FID under specific conditions or following treatments (e.g. antioxidants, inhibitors of cellular pathways). Figure 5. Open in a new tab Basic interpretation of FID experiment in arterioles using pressure myography to discern the primary endothelial-derived mediator. (A) Addition of inhibitor A results in a near complete loss of dilation in response to flow indicating that compound A is primarily responsible for FID in vessel #1. (B) In the presence of inhibitor B, vessel #2 dilates only to 40% of its maximal dilator capacity which represents ~50% of the vessel’s response to flow suggesting that mediator B is only partially responsible for FID. (C) A similar scenario is observed in the presence of inhibitor C for vessel #2. D) The combination of inhibitors B and C result in a near complete loss of dilation indicating that the residual dilation observed with the inhibitor of B is due to compound C. Assessing Endothelial-Independent Dilation: A key component during the assessment of FID in isolated vessels is determining whether a particular treatment that promotes a lack of response to flow (i.e. endothelial dysfunction) is due to a damaged endothelium versus a dysfunctional medial layer. A robust dilation to direct VSMC vasodilators should alleviate concerns of VSMC dysfunction despite a lackluster FID. As mentioned, the addition of papaverine at the end of each experiment may accomplish this task by observing >70% maximal dilation. If >70% dilation is not observed, one can compare concentration-response curves (EC 50 ) between control and treated vessels in response to direct VSMC dilators or NO-donors (e.g. sodium nitroprusside). These tests verify VSMC integrity and help determine whether reduced FID reflects true endothelial dysfunction or impaired medial responsiveness, rather than suggesting redundancy of independent vasodilatory pathways D. En Face Preparations Endothelial function can also be tested using en face preparations of blood vessels either in combination with other functional readouts (e.g., Ca 2+ signaling; see other sections), or as a standalone readout. En face refers to “on face” preparations of blood vessels with endothelium on the apical side. This is performed by using a pair of microdissection scissors to cut the blood vessel of interest longitudinally and pinning it to a silicon type material or coverslip, taping down the blood vessel edges, or securing it between glass coverslips. This technique is especially helpful when the need is to perform high resolution imaging of intact endothelium that may include cellular localization of proteins within the cell ( Fig. 6 ) ( 103 ), localization of proteins or organelles in proximity to each other, or where these structures in endothelium may align with cells or arterial anatomical hallmarks outside of the endothelium (e.g., myoendothelial junctions, see other sections). It can also be used to visualize the intact artery, i.e., how endothelium aligns with internal elastic lamina and smooth muscle in one Z-view (a visualization of the structures through the thickness of the tissue, captured as a single optical depth plane or a stack projected into one view). Figure 6. Open in a new tab En face visualization of EC in artery and vein. En face views of third-order mesenteric arteries (A) and vein (B) stained with claudin-5 (green) and DAPI (blue). Arrows indicate the direction of blood flow. Note the lack of directionality in veins due to the lack of flow creates a more cobblestone-type appearance of the endothelium. The scale bar in each image is 10 mm. Image taken with permission from ( 103 ). A key component to ensuring viable ECs in en face preparations is to limit the number of bends in the vessel wall during the preparation to ensure the inner walls do not touch. The quality of the microscissors is also important, and the longitudinal cut should be done rapidly so damage to endothelium is minimal along the edges. When prepared in this way, with physiological buffer, the en face preparation can be used to measure physiologically relevant outcomes [e.g., Ca 2+ responses ( 104 ) or lipid uptake ( 105 ); see other sections], or immunostaining. For preparations in which immunostaining is the desired outcome, en face preparations generally use 4% paraformaldehyde, however acetone/methanol can also be used generally for membrane-bound proteins. Veins can also be used for en face imaging ( 106 ), but this can be especially difficult due to their low structural rigidity and high compliance, meaning they will collapse, and the endothelium will stick. For staining veins, it is advantageous to maintain an open lumen, which can generally be done with a small amount of perfusion with pipettes on one end of the vein, while the other end can be cut with the scissors. A negative to en face preparations is the blood vessels must be cut, so the physiological pressure cannot be maintained and the ability to measure contraction or dilation of the vessel is lost. However, the VSMCs are still physiologically viable and thus will still contract if an agonist is applied. For this reason, myosin light chain kinase inhibitors such as ML-7 or ML-9 have been used in en face preparations to allow the signaling aspect of VSMC to proceed, but the physical motion inhibited. An alternative to en face preparations is to invert arteries, thereby exposing the endothelium on the outside for easy microelectrode access and optimized imaging. This can be done for many arterial beds, and these inverted arteries maintain normal endothelial vasodilation and can be mounted in wire myographs for simultaneous force measurements ( 107 )( 53 ) E. Endothelial Tube Preparation Conducted Vasodilation: The spread of vasodilation along microvascular networks was reported by August Krogh in 1922. In the microcirculation of the frog hindleg web, he observed that “ chemical stimulation may cause dilatation over a large area of the web” and concluded that this response “ points definitely to the existence of a network of nerve fibrils in which conduction can take place in any direction ” ( 108 ). Nearly a half-century later in the hamster cheek pouch, Duling and Berne reported that local stimulation of an arteriole with ACh initiated “propagated vasodilation” that spread bidirectionally along the vessel and was graded in distance and amplitude ( 109 ). At that time, it was concluded that “ It seems most probable that the transmission system resides in either the VSMC itself or in a perivascular nerve plexus ” ( 109 ). A decade later the obligatory role of the endothelium in vasodilation to ACh was demonstrated ( 2 ). Six years later, vasodilation to ACh was shown to be conducted from cell to cell along the arteriolar wall, independent of innervation, blood flow, or pressure ( 110 ). Although myoendothelial gap junctions in arterioles had been resolved at the ultrastructural level ( 111 ), there were “… too few studies of the physiological interactions between endothelial and smooth muscle cells in the intact arteriole to permit conclusions regarding the role of gap junctions ” ( 110 ). Myoendothelial Coupling: In isolated pressurized arterioles of the hamster cheek pouch, the rise in VSMC [Ca 2+ ] i during vasoconstriction was reported to drive Ca 2+ through myoendothelial junctions and into ECs, thereby initiating the synthesis of NO to modulate VSMC contraction ( 112 ). Faced with the elusive nature of EDH and growing evidence for a functional role of myoendothelial gap junctions ( 113 ), efforts centered on defining the nature of signaling between ECs and VSMCs in resistance vessels. In isolated pressurized arterioles from the hamster cremaster muscle, pharmacological inhibition of luminal IK Ca and SK Ca channels eliminated a secondary conducted response, suggesting that following VSMC stimulation, Ca 2+ signaling in the endothelium triggers hyperpolarization and initiates conducted vasodilation ( 114 ). In pressurized feed arteries from the hamster retractor muscle, dual simultaneous intracellular recording with dye labeling resolved the endothelium as the cellular pathway for conducted vasodilation ( 115 ) with direct electrical coupling between ECs and VSMCs through myoendothelial gap junctions ( 116 ). Complementary experiments in vivo resolved the endothelium as the cellular pathway for ascending (i.e., conducted) vasodilation in response to skeletal muscle contractions ( 117 ). Ensuing studies resolved “fast” and “slow” components of conducted vasodilation in response to ACh by triggering electromechanical relaxation of VSMCs along the vessel wall initiated by local IK Ca and SK Ca channel activation, with an ensuing 'wave' of Ca 2+ along the endothelium releasing autacoids to promote pharmacomechanical relaxation ( 118 )( 119 ). Additional experiments have provided insight into the dynamics of Ca 2+ and electrical signaling through myoendothelial microdomains involving connexin isoforms, K + and Ca 2+ channels, as well as IP 3 and its receptors to provide bidirectional feedback between VSMC:EC signaling during vasoconstriction and vasodilation ( 120 )( 121 )( 122 )( 123 ). For evaluation of the myoendothelial junction, refer to Section F . Endothelial Tube Preparation: The endothelial tube preparation was developed to define properties of electrical and Ca 2+ signaling intrinsic to native, intact microvascular endothelium, independent of VSMC, perivascular nerves, blood flow, or circulating vasoactive factors. While cell culture can provide valuable insight into signaling mechanisms, culture of ECs can alter phenotype, genotype, and metabolism ( 124 ). Thus, cultured ECs may not fully represent their behavior in the vessel wall. As an alternative strategy, preparation of intact native endothelium provides greater physiological relevance of experimental findings. The mouse superior epigastric artery (SEA) is well suited based on obtaining relatively long (≥2 mm) unbranched segments of a resistance artery (diameter <100 μm). Initial experiments focused on optimizing experimental conditions for maintaining the structural and functional integrity of freshly isolated endothelial tubes ( 125 ). While Ca 2+ responses to ACh were enhanced ~twofold at 32°C versus 24°C, the instability of endothelial tubes at 37°C precluded their study at standard body temperature ( Fig. 1 ) ( 126 ). At cooler temperatures, ECs remained functionally coupled to one another, able to transfer chemical and electrical signals from cell to cell through gap junctions ( 127 ). Representative key findings from endothelial tube preparations are summarized in Table 4 . Table 4: Representative Key Findings from Endothelial Tube Preparations ✓ Voltage-insensitive ion channels (SK Ca /IK Ca ) can tune electrical conduction along the endothelium through changes in membrane resistance (mouse SEA) (127). ✓ With aging, enhanced current loss through SK Ca /IK Ca channels impairs electrical conduction along the endothelium (mouse SEA) ( 129 ). ✓ Microvascular endothelium adapts to aging by reducing Ca 2+ influx during elevated oxidative stress. Protection from oxidative stress sustains endothelial integrity during aging (mouse SEA) ( 130 ). ✓ Lymphatic endothelial tubes are devoid of KCa activity. Instead, depolarization to ACh may promote the conduction of contraction waves during lymph propulsion (mouse popliteal) ( 137 ). ✓ L- and T-type voltage dependent Ca 2+ channels are not expressed in arteriolar endothelium; EC Ca 2+ responses to KCl or phenylephrine originate in VSMCs (rat cremaster muscle) ( 120 ). ✓ CGRP hyperpolarizes pulmonary artery ECs though activating K ATP channels (mouse lung) ( 128 ). ✓ Alzheimer’s Disease pathology represents a condition of altered K Ca and KIR channel function in posterior cerebral artery endothelium (mouse) ( 134 ). ✓ Acid sensitive ion channel 1a couples to IK Ca /SK Ca channels in mesenteric resistance arteries to mediate endothelium-dependent vasodilation (rat)( 132 ). ✓ Aging impairs endothelial KIR2 channel function in cerebral arterioles (mouse brain) ( 135 ). ✓ Aging increases the resilience of posterior cerebral arteries of males to oxidative stress while females are inherently protected (mouse) ( 136 ) Open in a new tab Microdissection, Digestion, and Trituration: Selected vessels are dissected from the surrounding tissue in physiological salt solution (PSS; in mM: 140 NaCl, 5 KCl, 2 CaCl 2 , 1 MgCl 2 , 10 HEPES, 10 glucose, pH 7.4). Please note that the PSS differs from the solution described in Table 2 , which is most commonly used for isometric preparations. This difference is due to variations in laboratory protocols. Subsequently, the vessels are cannulated onto a glass pipette [outer diameter, 50-80 μm; 1B100-4, World Precision Instruments (WPI), Sarasota, FL, USA] at one end. Residual blood is gently flushed from the vessel lumen. Vessels are cut into segments (length, 1-3 mm) and placed into PSS containing 0.62 mg·mL −1 papain, 1.0 mg·mL −1 dithioerythritol and 1.5 mg·mL −1 collagenase, and incubated for 30 min at 32°C ( 126 ). Given the variability in enzyme activity for each lot/batch, incubations may need to be adjusted to ensure sufficient breakdown of the extracellular matrix without compromising the integrity of the final preparation. Following partial vessel digestion, the enzyme solution is replaced with PSS lacking enzymes, and segments are transferred to a fresh vessel chamber mounted on a microscope stage platform for dissociation. VSMCs and adventitia are removed by gentle trituration though borosilicate glass capillary tubes that have been pulled (P-97, Sutter Instruments, Novato, CA, USA) and heat polished. To optimize trituration, the internal diameter of the capillary tube tip (~100 μm) should be ~10 μm larger than the diameter of the vessel segment. Following isolation, the endothelial tube is secured at each end using blunted heat polished micropipettes (diameter, 60-80 μm) positioned with micrometers at each end of the stage platform. The successful removal of VSMCs can be confirmed with visual inspection ( Fig. 1 ) and cellular survival confirmed with vital dyes ( 128 ). The endothelial tube preparation has been studied across multiple species from vascular beds including skeletal muscle ( 127 )( 129 )( 130 )( 131 ) mesenteric ( 132 )( 133 ), brain ( 134 )( 135 )( 136 ), and lung ( 128 ) as well as lymphatics ( 137 ). Note that the digestion protocol (above) requires modification to produce pulmonary endothelial tubes. Pulmonary arteries contain multiple elastic laminae to accommodate stretch associated with respiration ( 138 ) Therefore, to generate tubes of pulmonary endothelium, the addition of elastase (30 μg/mL) is required during the 30 min enzymatic digestion ( 128 ). Physiology: Ca 2+ and Electrical Signaling Intracellular Ca 2+ is a key regulator of endothelial function and endothelium-dependent regulation of vasomotor tone. Therefore, the study of Ca 2+ signals in native, intact endothelium is critical for understanding how ECs respond to physiological and pharmacological stimuli independent of influences of blood flow and circulating factors, surrounding VSMCs, tissue parenchyma, and associated nerves. This preparation is suitable for evaluation of Ca 2+ responses intrinsic to the endothelium ( 125 )( 130 )( 137 ) . A unique aspect of the endothelial tube preparation is the ability to evaluate the coordination of intracellular Ca 2+ signals among ECs ( 139 ). Electrical signaling is integral to endothelial function ( 115 , 116 )( 127 , 129 , 137 )( 128 )( 134 )( 135 ). Upon stimulation, the spread of hyperpolarization along the endothelium and through myoendothelial gap junctions to VSMCs leads to vasodilation ( 116 ). Native endothelial tubes provide a way to evaluate electrical conduction along the endothelium without influences of surrounding cells. By separating two microelectrodes at various distances (50-2000 μm) along an endothelial tube, the amplitude and decay of hyperpolarization and depolarization can be evaluated in response to current injection ( 127 ). The endothelial tube preparation also enables evaluation of Ca 2+ and electrical responses to agonists ( 120 )( 137 )( 128 )( 139 ). In addition to serving a source of energy, mitochondria are integral to endothelial signaling in regulating vascular tone ( 140 ). The endothelial tube provides a unique approach to evaluate the regulation of mitochondrial membrane potential ( 136 ). Oxidative stress is a hallmark of endothelial dysfunction, and the endothelial tube enables evaluation of endothelial ROS production ( 133 ). In addition, the endothelial tube can be utilized to evaluate cell viability and death under well-defined experimental conditions ( 130 ). Endothelial tubes have been utilized to evaluate factors such as sex, age, and disease( 130 )( 134 )( 136 ) ( 133 ). In addition to providing insight into physiology and pathophysiology, the endothelial tube preparation can be applied in genetically modified mice to determine how altering the presence of specific proteins affects the biophysical properties of microvascular and lymphatic endothelium. Nevertheless, a key limitation is that the endothelial tube preparation has yet to be cannulated and pressurized, thus it has so far been studied in the deflated (i.e., collapsed) state. In summary, the endothelial tube represents a versatile preparation with unique advantages for evaluating the biophysical properties of native intact vascular endothelium. It may thereby serve as a useful tool with broad applications for studying endothelial function. F. Myoendothelial Junction Visualization Myoendothelial junctions (MEJs; also called myoendothelial projections) are cellular extensions in the extracellular matrix that allow for extremely close physical proximity between ECs and VSMCs, reminiscent of neuromuscular junctions (thus the name). As described above, MEJs were first identified using transmission electron microscopy (TEM) by Rhodin in 1967 in terminal arterioles from rabbit ( 109 ). The cellular extensions composing MEJs are primarily, but not exclusively ( 141 ), of endothelial origin, depending upon the source of the blood vessels. The MEJs physically resemble cellular invadopodia, and depending on tissue bed, can be quite prevalent between ECs and VSMCs, especially as the arteries get smaller, with few to any MEJs in conduit arteries. In terms of size, MEJs can be ~0.5 μm in diameter and ~0.5 μm in length with up to 4-6 MEJs per single EC ( 142 ) ( Fig. 8 ). The expression of MEJs in veins and lymphatics is much less understood or observed, and the interaction between capillary endothelium and pericytes, especially in the brain, are only recently being explored ( 143 ), although these are not termed MEJs. Figure 8. Open in a new tab Myoendothelial junction visualization and comparison. Left, classical TEM view and identification of MEJs, with asterisks indicating the lumen of the artery. Center, en face view of artery demonstrating the “holes” in the internal elastic lamina between endothelium and smooth muscle where MEJs form. Right, comparison of the number of TEM-identified MEJs to the predicted number of MEJs assuming holes in the IEL are a proxy for the structures. The scale bar in the left image is 1 μm, center image is 15 μm. Images taken with permission from ( 142 ). Similar to invadopodia, MEJs are cellular microdomains where caveola and multiple signaling molecules reside and may be enriched. In addition, gap junctions have long been identified at MEJs connecting ECs and VSMCs ( 113 )( 144 ), but not every MEJ may physically connect between the two cells ( 142 ), and/or the gap junctions between the endothelium and VSMC may be closed ( 145 )( 146 )( 147 ). It is from this perspective, i.e., that the ECs signal to VSMCs via gap junctions at MEJs, that endothelial function as a component to the physical location of MEJs were first described. Although MEJs were hypothesized to be an important component of electrical conduction several decades prior ( 110 )( 148 ), the first work to demonstrate a strong likelihood of MEJ involvement was when it was demonstrated that an elevation in VSMC Ca 2+ caused an increase in endothelial Ca 2+ and release of NO. However, the microdomain nature of MEJs now make it a unique signaling hub of its own in endothelium, likely independent of what may be traversing gap junctions from VSMCs. For example, MEJs can have unique protein, enzyme, lipid, and ion channel distribution, and Ca 2+ signals that can arise from MEJs based on endothelial specific agonists ( 104 )( 142 )( 149 )( 150 ). Endothelial function pertaining to MEJs can be difficult to accurately assess, but en face preparations (described above) are the main mechanism in which this can be performed. In the past, observing MEJs was performed by TEM as this definitively identified the anatomical structure; however, this prevented physiological testing. Using en face preparations, the holes in the internal elastic lamina separating endothelium and VSMCs are now considered MEJs ( 142 ) ( Fig. 8 ), and so localization of proteins, as well as events such as Ca 2+ dynamics near these “holes” can be used as a proxy for cellular events occurring at MEJs. Using the en face preparations for MEJs, we are beginning to understand how this unique structure may contribute to overall EC function and pathology. G. EC Ca 2+ Imaging Increases in EC Ca 2+ activity, whether the events are focal (localized to a discrete location) or global (occurring uniformly across a cell or propagating from one location along the length of cell) are essential to a number of physiological processes including angiogenesis ( 151 )( 152 ), vascular permeability ( 153 ), leukocyte transmigration ( 154 ), and importantly for the current guidelines, modulating the magnitude of vasodilation ( 155 ). Our insight into the Ca 2+ dependence of vascular responses and our ability to discern the subtle characteristics of and underlying molecular contributors to specific Ca 2+ events all stem from the technological and experimental advances that have unfolded over the last four decades. The advent of chemical Ca 2+ indicators (CCIs) and genetically encoded Ca 2+ indicators (GECIs) enabled robust detection of intracellular Ca 2+ concentrations over time, providing insight into Ca 2+ dynamics. Advances in confocal and multiphoton microscopy further allowed imaging at sufficient speeds to resolve even the most discrete Ca 2+ signals. In the following sections, we will detail the methods of detection, discuss the range of imaging modalities, introduce analysis methods, and define commonly used terminology related to endothelial Ca 2+ events. Where relevant, we will put these concepts together in context of EC function in intact endothelial preparations described in these guidelines, and/or guidelines previously published in this journal ( 1 ). Methods of Endothelial Ca 2+ Detection: For these guidelines, we will refer to Ca 2+ influx as the movement of Ca 2+ ions from the extracellular space across the plasma membrane through Ca 2+ permeable channels. Further Ca 2+ release will refer to movement of Ca 2+ ions out of internal stores/sources, such as the endoplasmic reticulum, into the cytosolic space. Ca 2+ flux will be a generic term used to encompass any increase and/or decrease in the concentration of free Ca 2+ regardless of Ca 2+ source or cellular localization. Today there are two primary mechanisms of detecting Ca 2+ concentrations, namely CCIs and GECIs (see Table 5 for advantages and disadvantages of each). Table 5. Considerations for Intracellular Ca 2+ Detection Chemical Ca 2+ Indicator (CCI) Advantage Disadvantage ✓ Wide range of Ca 2+ affinities (low-high Kd values) ✓ Can buffer intracellular Ca 2+ ✓ Does not require transfection or permanent modification to the genome to image the CCI ✓ Difficult to obtain cell-type specificity when loading a CCI ✓ Over time the CCI can be extruded and/or can move to adjacent VSMCs via gap junctions. ✓ Difficult to load for in vivo preparations Genetically Encoded Ca 2+ Indicator (GECI) Advantage Disadvantage ✓ Allows for utilization of genetic elements to drive the GECI of interest in a specific cell-type ✓ Can buffer intracellular Ca 2+ ✓ GECI remains within the targeted compartment and is not extruded ✓ Requires transfection or permanent modification to the genome for expression of the GECI ✓ Genetically driven expression ideal for in vivo preparations Open in a new tab CCIs (which have been thoroughly reviewed elsewhere, see ( 156 ) are structurally similar to known Ca 2+ chelators/buffers but with a fluorophore allowing for detection of Ca 2+ concentrations. Many CCIs used for modern EC Ca 2+ studies contain an acetoxymethyl (AM) ester group, which improves the membrane permeability and, once de-esterified by intracellular esterases, maintains the indicator in the cell compartment of interest minimizing extrusion (although limited extrusion outside of these compartments still occurs). With CCIs, there are no standard loading protocols, rather they are fine-tuned for the preparation of interest and can include the addition of Pluronic-F127 and dimethyl sulfoxide (DMSO) to further improve membrane permeability and dispersal of the CCI within the loading solution. For EC studies, it is important that the loading buffer containing the CCI be directly exposed to the EC layer. Thus, in intact pressurized vessels the loading buffer must be introduced into the lumen and the excess then rinsed out. The CCI taken up by ECs is then equilibrated to allow for de-esterification. The length of time the EC layer is exposed to the CCI containing solution must be titrated to avoid loading the innermost layer of the vascular smooth muscle. For en face or EC tube preparations, where the ECs are fully accessible, exposing cells to the CCI in the bath solution is sufficient for loading, so long as the concentration and incubation time are optimized. Non-specific loading of the VSMCs is less of a concern in en face preparations, as the EC layer can be observed simply by adjusting the focal plane of acquisition. Since loading protocols vary with vessel type (artery, lymphatic, or vein), vessel bed (e.g., skeletal muscle arteries, cerebral arteries, etc.), animal model (e.g., rats, mice, etc.), experimental preparation (e.g., intact vessels, isolated ECs, en face ) and indicator (e.g., Fluo-4, Fura-2, Oregon Green BAPTA-488), we included representative references that provide a good starting point for those embarking on studies utilizing CCIs ( Table 6 ). Because murine arterial ECs do not always load effectively, ( 157 ) the development of GECIs has expanded this field significantly. Table 6. Commonly Used Indicators in the EC Literature CCI Experimental Preparation Reference Fura-2 ✓ pressurized rat cremasteric arterioles ( 158 ) ✓ pressurized hamster cremasteric arterioles ( 159 ) ✓ mouse EC tubes from inguinal efferent-axillary afferent lymphatics ( 137 ) ✓ mouse EC tubes from superior epigastric artery ( 125 ) Fura-PE3 ✓ pressured hamster cheek pouch arterioles ( 119 ) Fluo-3 ✓ pressured hamster cheek pouch arterioles ( 160 ) Fluo-4 ✓ en face mouse mesenteric arteries ( 122 ) ✓ pressurized hamster retractor feed arteries ( 123 ) ✓ mouse EC tubes from superior epigastric artery ( 126 ) ✓ rat EC tubes from cremasteric arterioles ( 120 ) Oregon Green 488 BAPTA-1 ✓ pressurized rat cremasteric arterioles and mesenteric arteries ( 161 ) Cal 520-AM /Calbryte ™ 520 ✓ small intramyocardial coronary arteries (IMCAs) human endothelial cells ( 162 )( 163 ) GECI Experimental Preparation Reference GCaMP2 ✓ Intravital mouse cremaster preparation ( 118 ) ✓ en face and pressurized mouse mesenteric arteries ( 122 )( 164 ) GCaMP6f ✓ pressurized mouse lymphatics ( 165 ) GCaMP2, GCaMP5, and GCaMP8 ✓ isolated mouse mesenteric arteries and veins, GCaMP8 in capillaries ( 166 ) Open in a new tab In the early 2000s, mouse models expressing GECIs were generated to allow for in vivo studies of dynamic cellular processes. A National Heart Lung and Blood Institute (NHLBI) Regional Resources Program (R24) funded consortium was developed known as the Cornell/National Heart Lung Blood Resource for Optogenetic Mouse Signaling (CHROMus), ( 167 )( 168 ) which became the go-to resource for obtaining indicator mice for the study of cardiovascular, lung, and blood disorders. These mice were generated using bacterial artificial chromosome (BAC) recombineering technology or using established promotor constructs, where the transgenic construct used for insertion contained the GECI downstream of a functional promoter for the cell-type of interest ( 168 )( 118 ). CHROMus developed ~25 strains of mice (now all commercially available from The Jackson Laboratory) using well-characterized and cell-type specific promoters which targeted ECs, VSMCs, cardiomyocytes, and neuronal cells ( 167 )( 169 ). One such CHROMus line, Cx40BAC-GCaMP2 (JAX Strain# 025404), which expresses GCaMP2 in arterial and venular ECs was the first GECI strain utilized in endothelial studies. GCaMP2 is a circular permutated GFP molecule flanked by Ca 2+ sensitive domains that stabilize the fluorophore when bound to Ca 2+ , allowing for detection of fluorescence ( 169 )( 170 ). Further, GCaMP2 was specifically designed to function optimally at physiological temperatures ( 171 ) which is why it has been used for many years for both in vivo ( 118 )( 172 )( 173 )( 174 ) and ex vivo ( 122 )( 175 ) preparations. In recent years, different GECIs have been developed that benefit from improved signal/noise ratios, reduced background fluorescence, and faster response kinetics. GCaMP5 ( 166 ), GCaMP6f ( 165 ), and GCaMP8 ( 166 )( 176 ) are all examples of newer GECIs that have been used for the study of arterial, venular, and/or lymphatic Ca 2+ signaling dynamics. In 2015, a seminal paper in the field of neuroscience was published that generated ~21 different mouse models which provided an alternative to the CHROMus strains, as expression of the GECI could be more tightly controlled simply by crossing to a cell-type specific driver strain (e.g. Cre-recombinase, tetracycline-controlled transactivator) ( 177 ). One such strain, Ai95(RCL-GCaMP6f)-D (JAX strain #028865), was generated by knocking in a GECI that contained an upstream floxed-STOP cassette which prevented transcription. This strain, when bred to a Cre-recombinase expressing mouse, removes the STOP cassette and allows for expression of the indicator in the desired cell-type of interest. Since 2015, this strain has been used experimentally for the study of lymphatic and arterial EC function ( 165 )( 157 ). The Ai95(RCL-GCaMP6f)-D strain, and others like it, will be useful for the future of the vascular field in general, as this strain can be crossed to mice that conditionally express Cre and possess a floxed allele for any gene of interest (with non-floxed littermates as the control). Thus, in this case GCaMP6f expression functions both as a ‘reporter’ of proper Cre-induction and as a Ca 2+ indicator, all while being compatible with loss of function studies of any gene of-interest in a cell-type specific manner. How CCIs and GECIs compare head-to-head still needs to be explored, as new indicators continue to be developed. That being said, Ledoux et. al ., compared the descriptive parameters (e.g., amplitude, area, frequency) of Fluo-4 and GCaMP2 in detecting spatially restricted Ca 2+ signaling events in mouse mesenteric arteries prepared en face and found them to be comparable ( 122 ). In short, both CCIs and GECIs can be used for studying EC Ca 2+ dynamics, but determining which one is the most appropriate for a given study depends on factors including the tissue/cell preparation used, animal model, cell specificity, and available imaging modalities. Methods for Imaging Ca 2+ : Since both CCIs and GECIs detect Ca 2+ via fluorophores and require high frequency image acquisition and spatial resolution, confocal ( 178 ) and multiphoton microscopy ( 161 )( 176 )( 179 ) are the primary techniques employed. A brief examination of the current literature demonstrates that both spinning disk ( 122 )( 165 )( 166 ) ( 157 )( 178 ) and laser scanning confocal microscopy ( 120 )( 161 ) are used to image EC Ca 2+ . Regardless of the exact model or type of microscopy used, some of the important considerations for imaging EC Ca 2+ are as follows: Practical tips: Utilize the lowest laser setting you require to image the Ca 2+ event of interest. High laser power can result in tissue damage and impact the longevity of the preparation. Minimize the amount of time that the preparation is exposed to the laser. Minimizing exposure time can be as easy as making sure the laser is only on to the preparation during an experimental run and/or when focusing. Utilize the Nyquist Theorem, also referred to as the sampling theorem, to determine the appropriate speed of acquisition ( 180 ). In the context of EC Ca 2+ , the sampling rate must be at least twice as fast as the duration of the fastest Ca 2+ event being recorded ( 181 ). If the Nyquist criterion is not met, aliasing can occur and/or Ca 2+ events may be missed completely. When imaging Ca 2+ signals, high magnification/high-numerical aperture (NA) objectives and/or a confocal system with zoom capabilities must be used. Too large a field of view could result in loss of resolution required for distinguishing discrete Ca 2+ events. Commonly used Terms for EC Ca 2+ Activity in Intact Structures: The first publication demonstrating EC Ca 2+ responses in intact arterioles was based on rat cremaster and published by Falcone, et. al . in this journal in 1993 using the ratiometric Ca 2+ indicator, Fura-2 ( 158 ). Although the image quality of the time did not allow for detection of focal events, this study was the first to show an increase in [Ca 2+ ] i from basal conditions following exposure to flow and the EC-dependent vasodilator ACh. Since then, terminology has emerged to more precisely describe different types of Ca 2+ signaling events. Indeed, the only way to truly differentiate events from one another is by the use of pharmacological agents (agonists and antagonists) and/or genetically modified mice to tease out the underlying mechanism coupled with sub-cellular localization of the Ca 2+ event. We have summarized some key terms related to EC Ca 2+ below and displayed these pictorially in Fig. 9 , however, it should be noted that several excellent reviews have been published on this topic in recent years ( 181 )( 182 )( 183 )( 184 )( 185 ). Figure 9. Open in a new tab Common Endothelial Cell (EC) Ca 2+ terminology. (A) EC-specific focal calcium events include IP3 receptor mediated calcium efflux events not within myoendothelial junctions (puffs), IP3 receptor mediated calcium efflux events within myoendothelial junction (pulsars) and calcium channel dependent influx events which can be within or outside the myoendothelial junction (sparklets). (B) Vascular smooth muscle cell (VSMC) initiated calcium events occur following stimulation of the VSMCs with phenylephrine, KCl, or a voltage-dependent calcium channel opener. Following the transfer of calcium and/or IP3 through gap junctions at the myoendothelial junction to the underlying endothelium, VECTors or Wavelets are observed. Ca 2+ waves (C) propagate along the length of the EC and (D) can also be transferred between adjacent ECs. Before expanding upon the intricacies of specific sub-populations of EC Ca 2+ events, it is worth noting some terms that can be used more generally. For instance, increases in [Ca 2+ ] can be referred to as ‘Ca 2+ signals’, ‘Ca 2+ activity’, and/or ‘Ca 2+ events’. Discrete, or spatiotemporally restricted, Ca 2+ signals are often referred to as ‘local’ or ‘focal’. Elementary Ca 2+ events are the most fundamental building blocks of Ca 2+ activity, such as the opening of individual Ca 2+ channels, which summate to result in larger Ca 2+ events. Intracellular Ca 2+ ‘waves’ propagate along the length of the EC via a Ca 2+ -induced Ca 2+ release mechanism (intracellular), while ‘global’ Ca 2+ increases are typically observed more uniformly across the entire cytosol of the cell. It should be noted that the term ‘Ca 2+ waves’ has also been used to describe intercellular Ca 2+ events that can propagate between adjacent ECs ( 119 )( 164 ). Thus, it is important when presenting Ca 2+ wave data to distinguish intracellular from intercellular events. Further, there are spontaneous Ca 2+ events occurring in the absence of a pharmacological stimulus, ( 122 )( 182 ) while agonist-induced Ca 2+ events, occur following exposure to a direct activator of a Ca 2+ channel or an activator of G-protein coupled receptors (e.g., G q ) that signal via Ca 2+ . It was observed that in cultured HeLa cells the presence of Ca 2+ blips, an elementary Ca 2+ signal originating from inositol 1,4,5-trisphophate (IP 3 )-dependent release of Ca 2+ from the endoplasmic reticulum, and Ca 2+ ‘puffs’, which are due to simultaneous activation of numerous blips ( 186 ). It was also observed that Ca 2+ ‘pulsars’ which were defined as spatially restricted IP 3 -dependent Ca 2+ signals within myoendothelial junctions ( 122 ). Specifically, the descriptive parameters between these endothelial Ca 2+ pulsars differed significantly from the more traditional Ca 2+ puffs observed in Xenopus oocytes and cultured cells ( 186 ) ( 187 )( 188 ). It is unsurprising that the characteristics of Ca 2+ events differ so dramatically depending on cell-type studied and sub-cellular location. Thus, it is worth distinguishing that IP 3 -dependent Ca 2+ events within myoendothelial junctions are ‘pulsars’ ( 122 ), while IP 3 -dependent Ca 2+ events occurring elsewhere in the EC are ‘blips’ or ‘puffs’ (dependent on the level of IP 3 receptor activation) ( 189 ). The subtle difference in nomenclature is important to recognize because only a sub-set of IP 3 -dependent Ca 2+ events (~70%), occur within myoendothelial junctions ( 161 ). To state that a Ca 2+ event is a ‘pulsar’, it needs to be shown that the event occurs within a hole in the internal elastic lamina (IEL) where myoendothelial junctions occur and is abolished in the presence of an IP 3 receptor blocker ( 120 )( 122 )( 161 )( 190 ). Experimentally, it is possible to determine the localization of a Ca 2+ event by simultaneously imaging the IEL. Although the IEL has some autofluorescence in the green spectrum, incubating the preparation with Alexafluor 633 hydrazide makes the holes in the IEL more apparent in the red spectrum which allows for dual imaging of Ca 2+ and the IEL in the green and red spectra, respectively ( 161 )( 191 ). Indeed, IEL autofluorescence can impede the detection of green fluorescence based GECIs in intact pressurized preparations, which is why they have been used most prominently with en face preparations where the curvature of the vessel is not a factor and the influence of the IEL can be minimized using confocal imaging. The first emergence of the term ‘sparklet’ was in reference to local EC Ca 2+ influx through transient receptor potential vanilloid 4 (TRPV4) channels, that occurred both within myoendothelial junctions and elsewhere within the cell ( 104 ). These TRPV4 sparklets were only observed in the absence of IP 3 -dependent Ca 2+ events, which were inhibited by incubating the preparation with either cyclopiazonic acid [to block sarcoplasmic reticulum/endoplasmic reticulum Ca 2+ -ATPase (SERCA)] or U73122 [to inhibit phospholipase C (PLC)] ( 104 ). Low intraluminal pressure was later found to be a physiological stimulus to increase TRPV4 sparklet induced Ca 2+ activity ( 161 ). Although initially used only in reference to TRPV4-mediated activity, the term ‘sparklets’ was adopted for any Ca 2+ influx event, with ‘sparklets’ being appended to the name of the associated Ca 2+ channel. Over the last decade transient receptor potential vanilloid 3 (TRPV3) sparklets ( 192 ), transient receptor potential A1 (TRPA1) sparklets ( 193 )( 157 ) and Ca 2+- permeable N-methyl-D-aspartate receptor (NMDAR) sparklets ( 194 ), have been observed in ECs from cerebral arteries. The physiological activators, and pharmacological activators/inhibitors to experimentally probe each of these “sparklets” has been well described in a recent review ( 185 ). Thus far, we have described Ca 2+ events that are stimulated by channels on the plasma membrane or within the compartments of ECs. In the following section we will focus on an additional sub-set of endothelial Ca 2+ events caused by activation of the overlying VSMCs and mediated by heterocellular communication via gap junctions at the myoendothelial junction. It has been shown that activation of VSMCs with the α 1 adrenergic agonist, phenylephrine, or high concentrations of KCl resulted in increased EC Ca 2+ activity ( 160 ). This VSMC-dependent agonist-induced increase in EC Ca 2+ activity was attenuated in the presence of nifedipine, suggesting a role for voltage-dependent Ca 2+ channels (VDCC) in the response. The activation of VSMCs was proposed to increase second messengers (Ca 2+ and IP 3 ) which then diffuse through gap junctions at the myoendothelial junction to activate Ca 2+ signaling events in the endothelium. It was proposed that any vasoconstrictor that can increase Ca 2+ and IP 3 can trigger this feedback loop, which could play an important role in controlling vessel diameter and the modulation of blood flow by attenuating VSMC contraction ( 160 ). Following improvements in imaging technology, Ca 2+ ‘wavelets’, which were spatiotemporally restricted IP 3 receptor-mediated EC Ca 2+ events following phenylephrine exposure, were first described and supported the idea that IP 3 was a key second messenger ( 123 ). Several years later, VDCC-dependent EC Ca 2+ transients (termed VECTors) were described which were endothelial Ca 2+ events triggered following direct activation of L-type VDCCs on VSMCs using the L-type Ca 2+ channel agonist, Bay K8644 ( 120 ). The presence of VECTors and wavelets collectively support the idea that Ca 2+ and IP 3 are second messengers that can mediate feedback between adjacent cell layers. Furthermore, when the connections between VSMCs and ECs were eliminated by isolating the endothelium (see endothelial tube preparation section above), direct exposure ECs to phenylephrine, KCl, and Bay K8644 did not increase Ca 2+ signaling ( 120 ). Thus, when studying Ca 2+ events that rely on EC: VSMC communication, the endothelial tube preparation can be a powerful experimental control for eliminating any role for VSMCs. It should also be recognized that when ECs are stimulated and their [Ca 2+ ] i increases, the [Ca 2+ ] i in surrounding VSMC can decrease ( 112 ) In such manner, reciprocal Ca 2+ signaling between ECs and VSMCs may contribute to vasomotion ( 195 ). To date, there has been a range of EC Ca 2+ events defined within the literature. When embarking on new studies in this area it is important to consider a few key points ( Table 7 ): Table 7: Key Points for EC Calcium Activity in Intact Structures: ✓ What type of Ca 2+ signal are you investigating, and does it occur in a spatially restricted manner? Do you need a fast indicator or not? ✓ Are you able to simultaneously image Ca 2+ and the IEL to determine if events are happening within potential myoendothelial junctions? Lymphatic vessels do not have as robust an IEL layer, so simultaneous imaging of Ca 2+ and the IEL may be more important in arterial and venular preparations. ✓ Are you using the appropriate CCI or GECI to image Ca 2+ events within the cellular compartment of interest, and with the suitable kinetics to detect the full dynamic range of the response? ✓ Are you imaging fast enough and do you have a sensitive enough detection system to image fast and/or discrete events? ✓ Are you using the appropriate pharmacological agents to activate and/or inhibit the channels involved in the responses being studied? Do genetically modified mice exist that could be used to validate the pharmacological approach? ✓ Are you using the appropriate in vivo (intravital preparations) or ex vivo (pressure myography, wire myography, en face) vascular technique to image the Ca 2+ events you are studying. For instance, if pressure is an important variable, an en face preparation would not be appropriate. Open in a new tab Analyzing Ca 2+ Data: There is not one standard method or program used to analyze EC Ca 2+ events. Indeed, one of the challenges with pressurized vessels or in vivo preparations is the potential tissue movement ( 196 ), which makes automated analysis techniques difficult to implement. Despite this, researchers in this area have used commercially available programs ( 120 ), ImageJ plug-ins ( 165 )( 197 ), and custom software ( 104 )( 122 )( 173 ) ( 196 )( 198 ) to obtain vital descriptive characteristics of EC Ca 2+ events. In en face preparations and in cultured cells, where tissue movement is minimal, there has been forward progress in fully automated analysis ( 196 ), but the application of these technologies to dynamic vascular preparations requires further development. See section 4 Software Acquisition and Analysis for more details. In the early days of imaging endothelial Ca 2+ , an entire field of cells was analyzed as a singular unit, generating a single output regardless of the number of cells in the field of view (FOV) ( 158 ). This analysis evolved into examining Ca 2+ events within a FOV, across an individual cell, or to a focal region of interest (ROI) within a cell/FOV where discrete events occur. Instead of focusing on the wide array of analysis programs available, in this section we will highlight some of the key considerations for analyzing EC Ca 2+ events. For more details, please see the section below titled: “ Software for Endothelial Ca 2+ Imaging Analysis ”. What descriptive characteristics are used to define distinct Ca 2+ events? The array of descriptive parameters used to describe Ca 2+ events can differ between publications, but some of the most common are frequency of events, amplitude of response, duration of response, and area of response. Depending on the method of image acquisition, either the percentage of cells active per FOV (if distinct cell borders are visible, used most often in pressurized vessels where curvature of the vessel can impact the number of cells visible) or number of active sites per FOV (if a set FOV size is used routinely, more common in en face preparations) can be used to determine the relative activity level of a preparation. Is the event spontaneously occurring or does it require a physiological/pharmacological stimulus? When analyzing a stimulus-induced increase in Ca 2+ activity, the responses are usually compared to basal activity. Thus, it is not uncommon to record the activity at baseline and following a stimulus and to repeat these recordings in the presence of pharmacological antagonists to decipher the influx and/or release pathways underlying the Ca 2+ response. Ideally, a single FOV is used for collecting all the data described above so the activity of specific sites can be monitored longitudinally throughout the experiment. Is the Ca 2+ event focal? It has been reported that placing a ROI around an entire cell will often mask the focal events occurring within the cell ( 161 ) ( Fig. 10 ). Thus, it is important that smaller ROIs be utilized when discerning focal events, with data acquired at appropriate resolution. Of note, ROI size and manual placement of the ROI can be time consuming and requires fully and systematically scanning across each cell within the field of view; therefore, automated detection and quantification approaches using dynamic ROIs defined by the maximal area of the Ca 2+ event may provide an alternative strategy to reduce bias. Please see section “ 4. SOFTWARE ACQUISITION & ANALYSIS ” for further information. Does the Ca 2+ event propagate? To study intracellular Ca 2+ events, numerous ROIs need to be placed along the length of the cell to demonstrate the propagation of the wave and to determine propagation velocity. To study intercellular waves, Ca 2+ activity in adjacent cells needs to be studied with a remote cell being used as a control ( 139 ). Does your preparation have vasomotor activity? Vasomotor activity does not preclude the study of Ca 2+ activity. Indeed, when studying pressure or flow-dependent responses, utilizing intact vessels that will have tissue movement is an inevitable challenge. The key is to minimize adjustments to focus during the recording you plan to analyze. This requires a quick adjustment to focus before starting the recording and minimizing the adjustment throughout the recording. Ideally for determining frequency (number of events per minute) an entire minute is recorded with no tissue movement. Some groups, particularly those studying lymphatic vessels which display rhythmic contractile activity, inhibit contractile activity ( 165 ), when appropriate. Figure 10. Open in a new tab Quantification of Focal Ca 2+ Events Requires Sub-cellular Regions of Interest (ROI). (A) Ca 2+ activity was measured using an Olympus FV1000 laser scanning confocal microscope via the chemical calcium indicator Oregon Green 488 BAPTA-AM in isolated, pressurized rat cremasteric arterioles at 5 mmHg intraluminal pressure. ROIs were drawn around three cells (red borders and associated red trace, a-c) and subcellular ROIs were placed at sites of focal activity (blue boxes and associated blue trace). (B) Although whole cell ROIs can sometimes detect small sub-cellular changes in calcium activity (upward black arrow), in other instances, subcellular calcium events are not clearly detectable when using a whole cell ROI (red downward arrow). These data demonstrate that small ROIs are necessary to detect and quantify sub-cellular Ca 2+ events. Scale bar, 10 μm. Acquisition rate ~3Hz. Image taken with permission from ( 161 ). In summary, although there can be many iterations of techniques employed for the study of EC Ca 2+ signaling dynamics, there are some common considerations. The intention here is to provide an overview so that those new to the field understand the historical context of Ca 2+ imaging and have a greater appreciation of important experimental criteria when embarking on a new study. H. Endothelial Barrier Integrity Microvascular exchange between the blood and tissues occurs primarily in the capillaries and postcapillary venules. However microvascular leakage of plasma components becomes elevated under inflammatory conditions, and when severe can lead to swelling, poor delivery of oxygen to cells, and tissue dysfunction. As described above, ECs form the main blood-tissue barrier, with subcellular components as the glycocalyx surface layer, tight and adherents junctions, focal adhesions, and vesicular transport mechanisms all contributing to endothelial barrier and transport functions ( 199 )( 200 ). The permeability of the endothelium to a specific solute provides important insights into barrier integrity. Permeability (Ps) describes the properties that allow solutes to cross the endothelium and is often derived from experimental adaptations of Fick’s principles. Fick’s first law strictly applies to diffusion, and the diffusion coefficient is a material constant. The permeability incorporates additional factors including barrier thickness and partitioning of the solute into the membrane. It is important to note that flux is defined as transport rate per unit area. Solute transport in vivo is influenced by variables such as capillary perfusion, solute size, and charge, and potentially by pressure-driven filtration. Tracer-based imaging techniques are commonly used to estimate permeability, with the assumption that these tracers behave similarly to endogenous molecules ( 200 )( 201 ). J s = P s ( C P L - C T ) where J s is the solute flux, P s is the permeability to the solute, S is the surface area, C P L is the solute concentration in the perfusate/lumen, and C T is the solute concentration in the tissue. Dye Accumulation Assays: One of the simplest methods to determine the level of edema is to compare wet-to-dry weights of organs between a treatment group and control group. Related to this, a simple approach to assess microvascular leakage at the organ level is to measure the accumulation of intravenously administered tracer dyes or fluorophore-conjugated molecules in small rodents like mice. Typically, one or more tracers is administered via the tail vein or by retro-orbital injection, followed by a set duration of time and then whole-body perfusion to remove the tracers from the bloodstream. The organs are then harvested, weighed, and then tracer levels are measured either by imaging or fluorometry of whole-organ homogenates ( 202 , 203 )( 204 ). One option is also to measure dye accumulation on the skin ( 205 ). The advantages of these approaches are simplicity, the ability to measure leakage in multiple organs, the ability to use multiple tracers of different sizes and wavelengths to evaluate size-selectivity in a single animal, and the potential capability to perform the experiment with several mice on the same day. The disadvantages are that the endpoint is a crude measurement of leakage, only one time point can be evaluated, and mechanistic insights may be limited ( 200 ). Intravital Microscopy: Imaging of the microcirculation using fluorescence microscopy enables the study of intravenously injected tracers and their escape into the surrounding tissues over time. This approach is typically performed with anesthetized rodents and involves surgical procedures to expose a tissue for microscopy imaging. Thin tissues are most often used, including but not limited to the mesentery, cremaster muscle, and hamster cheek pouch. The quantification of microvascular leakage is done through image analysis. One option is to count how many visible leakage sites appear at planned time points in a predetermined viewing window ( 206 )( 207 ). A more common approach is to measure the integrated optical intensity (IOI) of the tracer in the tissues surrounding the postcapillary venules ( 208 )( 209 ). When multiple time points are measured over time, a continuous infusion of the tracer after the initial bolus is ideal to maintain a plasma steady-state for accurate assessment of microvascular leakage due to changes in permeability over time ( 210 )( 211 ). However, some models like those featuring hemorrhagic shock have massive systemic microvascular leakage of tracers compared to controls, making it difficult to maintain steady-state tracer levels in the plasma. In this case, the IOI of extravascular tissue is normalized to the intensity inside vessels ( 212 )( 213 )( 214 )( 215 ). Also, it is important to monitor arteriolar diameter to account for potential increases in IOI due to filtration rather than elevated microvascular permeability ( 200 ). An example of images and IOI measurements from a study of the impact of hemorrhagic shock and resuscitation on mesenteric microvascular leakage is shown in Fig. 11 . Advantages of intravital microscopy include real-time in vivo observations, the ability to observe dynamic processes, and the ability to directly view microvascular leakage sites and make localized estimations of changes in permeability that may be more dramatic than whole-organ measurements. Disadvantages include the need to anesthetize animals, which might have an impact on microvascular permeability, limitations in the types of tissues that can be observed, and potential inflammatory impacts of the surgical preparations needed ( 200 ). Figure 11. Open in a new tab Example of intravital imaging of the rat mesenteric microcirculation in a study of the impact of hemorrhagic shock and resuscitation (HSR) and treatment with intravenous sphingosine-1-phosphate (S1P) on microvascular leakage. On the left are representative fluorescent images from sham treatment, HSR, and HSR + S1P groups. The arrows point at “hot spots” of albumin leakage. The circles indicate areas with apparent extravascular albumin leakage. The graph on the right shows quantification of FITC-albumin extravasation. Integrated optical intensity (IOI) was measured in the extraluminal space adjacent to postcapillary venules. **p < 0.01, *p < 0.05 (one-way ANOVA followed by Tukey’s multiple comparisons test). Image taken with permission from ( 212 ). Single-Perfused and Isolated Microvessel Models: Intravital microscopy can also be used in conjunction with cannulation of capillaries or venules to determine P s (solute permeability) or Lp (hydraulic conductivity) in individual microvessels. This is particularly useful for mechanistic studies and understanding how the contents of perfusion solutions impact microvascular transport. Because the vessels are very small, glass micropipettes are used as cannulas and are guided using micromanipulators ( 216 )( 217 ). A second approach is to dissect and isolate venules, which are then transferred to a chamber bath and cannulated on both ends ( 218 )( 219 , 220 )( 221 ). For both approaches, the ability to rapidly change between a physiological perfusion solution containing tracer such as fluorophore-labeled albumin and no tracer (unlabeled albumin) is key for controlling the initial concentration difference across the endothelial wall ( C PL - C T ). This can be achieved in single perfused microvessels by using two pipettes to cannulate at a bifurcation ( 216 ) or in isolated venules using concentric pipettes (pipette within pipette method) or using a theta pipette ( 218 ) (type of double-barrel glass micropipette). To determine permeability, the tracer is rapidly introduced into the microvessel, and images are rapidly acquired over a short period of time to capture the leakage of the tracer into the surrounding tissue. A pre-determined box-shaped region of interest (ROI) around and including the vessel is used for analysis ( Fig. 12 ). The initial step increase in overall intensity within the ROI when the tracer is rapidly introduced ( Δ I f 0 ) divided by the volume of the capillary or venule ( V ) represents the initial ( C PL - C T ). The subsequent rise in overall intensity over time ( d I t ∕ d t ) 0 represents the flux of the tracer ( J s ). With the assumption that the microvessel’s geometry is cylindrical, its volume divided by the surface area (S) reduces to r/2 ( 216 ). With these components, the permeability coefficient can be derived from Fick’s first law of diffusion, using an approach adapted for our endothelial cell assays, resulting in the following expression: P s = ( 1 Δ I f ) ( d I f d t ) 0 ( r 2 ) Figure 12. Open in a new tab Determination of permeability in isolated venules. After a venule is dissected and excised, it is mounted on two glass micropipettes and bathed with 37 °C albumin-physiological saline solution (APSS) as shown in panel (A). The heights of the inflow and outflow reservoirs are used to control pressure and flow. (B) The fluorescence intensity of a window containing the venule and nearby extraluminal area is measured. The inflow reservoir is quickly switched from APSS without tracer to APSS containing a fluorochrome-labeled tracer such as FITC-albumin. This causes a step increase fluorescence intensity Δ I f 0 within the window. The Δ I f 0 is proportional to the number of the tracer molecules that have just entered the lumen. Immediately after this, the intensity increases gradually (dIf/dt)0, and this rate is proportional to solute transport across the microvascular wall. The intensity returns to the basal level when the tracer is washed out from the lumen. These measurements are then used to determine the permeability coefficient with the equation indicated in the narrative. Image taken with permission from ( 200 ). Another single-perfused microvessel model worth mentioning, but less commonly used now, is the determination of Lp with the modified Landis method, detailed elsewhere ( 222 ). For all the single-perfused and isolated microvessel models, the perfusion pressure can be tightly controlled allowing for precise determination of how other experimental factors affect diffusive permeability. Another advantage of the isolated venule approach is that factors from the tissues are absent and the bath solution can be tightly controlled to mimic conditions to be studied, such as the addition of particular inflammatory cells or mediators ( 220 )( 223 )( 224 )( 225 ). In addition, with isolated vessels, the absence of non-vascular cells simplifies the study of signal transduction pathways ( 226 ). The main disadvantage of these models is that they are relatively difficult to perform, requiring extensive training and time to obtain reliable results. Another concern with the isolated venule approach is potential damage to the vessels during isolation. However, when these types of models are optimized and produce baseline permeability coefficients are in the expected physiological range, they can be very powerful tools for gaining mechanistic insight about how endothelial permeability is controlled. In Vitro Endothelial Monolayer Assays for Evaluating Endothelial Barrier Function: Please refer to the next section for detailed instructions on culturing ECs. This section focuses solely on evaluating endothelial barrier function. Cultured EC models have become widely used for evaluating endothelial barrier function. Common diffusion-based models rely upon cells grown on a membrane that is permeable to solutes such as albumin or large dextrans. A second popular approach is to measure the electrical resistance of endothelial monolayers, either across cells grown on a membrane or on a measuring electrode. Both types of assays require several considerations. First is the type of cells to be used. Human umbilical vein ECs are easy to grow and commonly used, but in some cases ECs of microvascular origin may be more appropriate ( 212 )( 227 )( 228 )( 229 )( 223 ) ( 230 )( 231 )( 232 )( 233 )( 234 )( 235 ). Culture conditions are also very important. The membrane should ideally be coated with a matrix such as gelatin, fibronectin, or other basement membrane protein mixture that promotes sufficient focal adhesion. Seeding at a density that immediately achieves confluence and allowing 5-7 days for junctions to mature is also an important consideration ( 229 )( 234 ). For diffusion-based models, the choice of tracer is important. Fluorophore-conjugated albumin is typically chosen to model plasma albumin. A 1:1 ratio of fluorophore-to-albumin ratio is desirable to obtain a linear standard curve that reflects the protein concentration. Some FITC-albumin products have higher ratios making them brighter, but also have the risk of dissociation of FITC, which could make the apparent P s higher than the true P s . It is also worth noting that FITC can significantly alter the size and charge of albumin, affecting FITC-albumin flux measurements ( 236 ). Newer AlexaFluor fluorophores, which are brighter and have 1:1 labeling with albumin may be a better choice. Dextrans, labeled 1:1 with fluorophore, available in several molecular weight ranges, are another alternative and allow for determining permeability of different sized molecules ( 230 )( 237 ). One final consideration is about the cultureware limitations and whether the protocol will provide a single P s per experimental group, or multiple P s values from a single EC monolayer before and after an experimental intervention. For the single P s approach, an advantage is that cultureware that requires a relatively small amount of media and tracer can be used ( Fig. 13A ). In contrast, for multiple measurements of solute flux to obtain P s values before and after addition experimental interventions require a specialized chamber system that requires larger amounts of media [ Fig. 13B ( 238 )]. Figure 13. Open in a new tab Configurations of diffusion-based endothelial monolayer permeability assays. A. Transwell assay in which the apical side of the endothelial monolayer faces the upper chamber, representing the vessel lumen, and the bottom chamber represents the abluminal side. The cultureware is placed in an incubator for a set amount of time, then the measurements are made upon retrieval. Typically, only one measurement can be made due to the small volumes, and the impact removing the cells from the incubator can have on permeability. B. Endothelial monolayers grown on Snapwell inserts are placed in an Ussing chamber system, which has larger volumes of media in each chamber (5 ml each). The chambers are placed in a heated manifold and media bubbled with 5% CO 2 so that experiments can be run for long periods of time. Samples can be drawn over time, before and after treatment with inflammatory mediators or other compounds. Image taken with permission from ( 238 ). Measurement of transendothelial electrical resistance (TER) is another method commonly used in vitro to evaluate endothelial barrier function. One approach is to grow endothelial monolayers of cells with a membrane placed between chambers filled with media and place an electrode in each chamber. Early studies with this approach yielded resistance values in the range of 6.1 – 69 Ω·cm 2 for peripheral EC ( 239 )( 240 )( 241 ) ( 242 )( 243 ), and 160-800 Ω·cm 2 for brain EC ( 244 )( 245 )( 246 ). A second approach named electric cell-substrate impedance sensing (ECIS) has been to grow the cells on a small gold recording electrode, with a much larger counter electrode to measure TER ( 247 ). The general protocol utilizes a 1-V, 4000-Hz AC signal supplied through a 1-MΩ resistor, which approximates a constant-current source to obtain TER values that are typically 100-1000 times higher than with the two-chamber method, and has the advantage of better detecting changes in TER. In addition, there is a mathematical model that allows ECIS users to resolve relative contributions of intercellular versus focal adhesion ( 248 )( 249 )( 250 )( 251 )( 249 ). ECIS has the advantage of being easy to use and having high sensitivity to detect endothelial cell micromotion. However, ECIS results may not align with change is permeability for larger solutes, so interpretations should be made with caution. I. Isolated Lymphatic Vessels Studies Lymph Flow and Transport: As part of the circulatory system, the lymphatic system plays a critical role in maintaining fluid homeostasis (i.e., by removing excess interstitial fluid, along with all its components), providing a main trafficking route for immune cells, transport of cytokines ( 252 ) and incretins ( 253 ), and lipid absorption and transport. In most tissue beds, interstitial fluid steady states are achieved with a low-level filtration matched by an equivalent lymphatic outflow ( 254 ) resulting in the daily transport of 3-5 liters of lymph in humans through the thoracic duct on its return to the blood circulation. Lymph flow within a tissue can also vary dramatically (~10-fold) given the hydration status, vascular, blood perfusion to the specific tissue bed, and in certain cases influx of fluid to the tissue across an epithelial barrier such as in the mesentery ( 255 ). Lymphatic vessel adaptation to differences in regional steady state demands ( 256 ) along with likely regional differences in cell precursors result in different lymphatic collecting vessel contractile activity ( 257 ) broadly defined into either “pump” or “conduit” vessel behavior ( 258 ), which is primarily dependent on signaling by the lymphatic endothelium. Despite the significantly lower flow and shear stress present in lymphatics vessels as compared to the blood vasculature, lymphatic endothelial cells still can play an active role in contractile modulation through an exquisite shear sensitivity ( 259 ). In certain lymphatic vessels, such as the rat thoracic duct, an imposed flow by a pressure differential of just 5 cmH 2 O will cause a near complete cessation of spontaneous contractions and vessel dilation, termed flow-induced inhibition ( 260 ). However, a similar imposed flow with a pressure differential of 5 cmH 2 O in rat mesenteric lymphatic vessels may have a variable or only partial flow induced inhibition ( 260 ) highlighting the regional heterogeneity of the lymphatic contractile regulation ( 257 )( 261 ). Additionally, lymph transport would not be possible without the other major functions of the lymphatic endothelium, namely the intraluminal one-way check valves that restrict backflow and tight junctions between lymphatic endothelial cells (LECs) that regulate lymphatic permeability. Physiological Saline Buffers in Lymphatic Collecting Vessel Research: Isolated lymphatic vessel studies have used a range of buffering molecules and physiological saline solutions (PSS) of varying compositions over the past decades. While this section describes the commonly used buffer systems in lymphatic research, the authors strongly recommend taking into account the significant impact of buffer composition ( 55 ) and the necessary considerations regarding alternative buffers or supplemented components as discussed in the early section of these guidelines. Because the study of lymphatic vessels is technically challenging and limited to a small group of researchers, it is important to note that some publications have reported adaptations to the solution for lymphatic tissue. While these adaptations may not represent a consensus among the present authors, they have been successfully applied in specific contexts. Classically, isolated lymphatic vessel perfusion baths were bicarbonate buffered solutions aerated with 95% O 2 - 5% CO 2 , including the seminal work on lymphatic pacemaking by Dirk Van Helden ( 262 ). However, 95% O 2 is supraphysiological for arterioles and even less physiological for lymphatic vessels that have rather low partial pressures of oxygen as most lymph is both hypoxic and hypercapnic compared to arterial blood. pO 2 levels of 20-30 mmHg are recorded in peripheral lymphatic vessels ( 263 )( 264 ) and values as low as 6 mmHg ( 264 ) have been recorded in lymphatic collecting vessels. In the centrally located thoracic duct, pO 2 values were recorded between 30-60 mmHg ( 265 )( 266 ). The comparison between recorded O 2 and CO 2 pressures in vivo and applied levels in ex vivo experimental studies is further complicated by transmural differences in gas tensions between lymph and interstitium and the presence of vaso vasorum, at least in the larger collecting vessels ( 267 ), which may cause local deviations in oxygenation relative to the bulk lymph. Irrespective, 20% O 2 - 5% CO 2 bubbled bicarbonate solutions remain instrumental in many of the foundational electrophysiology studies on human lymphatic collecting vessels using a wire myograph approach ( 268 ) ( 269 ) ( 270 )( 271 )( 272 ). Regardless of the pO 2 , bicarbonate is a critical molecule for physiological pH regulation and as a substrate for various anion transporters with a documented effect on arterial tone ( 55 )( 272 ). It is worth noting that there is a paucity of studies utilizing isolated lymphatic collecting vessels and imposed flow gradients to assess lymphatic endothelial flow-mediated inhibition of contractions. The limited number of isolated lymphatic collecting vessel studies is further complicated by the differences in model organisms (specifically rats vs. mice), specific lymphatic collector or duct being studied, and the specific buffer composition (only a few of which are detailed below) used across labs. There has yet to be a detailed direct comparison of either lymphatic myogenic or endothelial function across the historically employed buffers. Our recommendation for future studies is to first utilize the buffer detailed in Table 2 when conducting their isolated lymphatic vessel studies to assess lymphatic endothelial function or flow-mediated inhibition of lymphatic contraction prior to pursuing the buffers below. An albumin supplemented PSS (APSS) buffer containing (mm: 145.00 NaCl, 4.7 KCl, 2.0 CaCl 2 , 1.17 MgSO 4 , 1.2 NaH 2 PO4, 5.0 dextrose, 2.0 sodium pyruvate, 0.02 EDTA, 3.0 (3-(N-morpholino) propanesulfonic acid (MOPS) and 10 g L−1 bovine serum albumin) that uses MOPS for pH buffering, in lieu of bicarbonate and CO 2 aeration due to the presence of albumin in the bath, has been used with success to assess lymphatic endothelial-mediated inhibition of contractions. The problem with gassing a protein containing solution can be circumvented by using a semipermeable membrane as described in this study ( 273 ).Rat lymphatic collecting vessels and ducts studied in APSS provided some of the first reports of flow mediated inhibition of contractions in the lymphatic vasculature ( 260 ). Recent contractile studies utilizing lymphatic collecting vessels incorporated 3 mM bicarbonate into a modified HEPES-buffered Krebs solution ( 274 ) ( 275 ). Although this solution provides a minimal level of substrate for ion transporters, the unphysiologically low bicarbonate concentration and absence of balancing CO2 bubbling make this solution inappropriate for maintaining physiological pH and this approach is not recommended. A direct comparison of the mouse inguinal axillary lymphatic contractile function was assessed in both this HEPES-Krebs PSS and a bubbled 21% O 2 - 5% CO 2 bicarbonate solution): 130 NaCl, 5.4 KCl, 0.5 mM NaH 2 PO 4 , 0.8 mM MgSO 4 , 22 mM NaHCO 3 , 5.5 Glucose, 1.8 mM CaCl 2 , as listed in ( 1 )( 58 ) and the above. There was a slight reduction in frequency noted in the bicarbonate buffered group ( 276 ) though whether that was due to slight differences in the ionic composition, the more complete regulation of cellular pH and anion transport balance by bicarbonate ( 58 ), or the elevated bath pO 2 was not further explored ( 276 ). It is worth noting that flow-mediated inhibition of contraction has yet to be published in isolated mouse lymphatic collecting vessels and the majority of isolated vessel studies assessing flow mediated inhibition of contractions have been performed with rat lymphatic collecting vessels. This includes two recent reports ( 277 ) ( 278 ) one of which used the APSS buffer while the other used the HEPES-buffered Krebs solution detailed above. Addition of 0.5% BSA helps stabilize proteins and the glycocalyx ( 279 ), minimizing protein denaturing and tissue decay during dissection and vessel cannulation. Solutions are sterile filtered and stable when stored at 4°C. The BSA supplemented Krebs is commonly used in the intraluminal solution to help support the lymphatic endothelial integrity, however BSA is typically omitted from the bath due to its drug binding ability. Given the extensive immune cell density on lymphatic vessels a high purity BSA is often required to prevent activation of inflammatory pathways by contaminants. There is a significant presence of macrophages along the lymphatic collecting vessels which can express arginase and reduce arginine bioavailability and NO production( 280 ). Experiments testing flow should consider the use of supplemented arginine for NO dependent studies ( 277 ) Microdissection of Lymphatic Vessels and Pressure Myography: Research using isolated lymphatic collecting vessels first started in large mammals (typically bovine mesenteric vessels) owing to the difficulty in observing and isolating the nearly transparent (when free of chyle) lymphatic collecting vessels and the abundance of lymphatic collecting vessels draining the mesentery ( 281 )( 282 )( 283 )( 284 ) ( 285 , 286 ). The collecting lymphatic vessels and large collecting ducts are the primary lymphatic vessels used for studies and are characterized by having a continuous basement membrane and coverage with a specialized type of smooth muscle cells, i.e., lymphatic muscle cells (LMCs), which display unique electrical pacemaking activity that entrains their coordinated contraction ( 165 ) ( 261 ). However, the isolated lymphatic vessel field eventually transitioned into using rodents, primarily rats and guinea pigs, which can readily be housed and maintained compared to ovine or bovine models. Conspicuously absent were comparable lymphatic collecting vessels studies from mice, aside from a few studies using DDY mice ( 287 ), due to significant differences in muscle cell investiture and contractility of lymphatic collecting vessels located in the thoracic or peritoneal cavity such as the mesenteric and thoracic duct. A landmark study in 2011 demonstrated that the mouse popliteal lymphatic collecting vessel had robust contractions that could be inhibited by NO ( 288 ) which led to the identification of various collecting lymphatic vessels located in the periphery, and not the visceral cavity, that had robust and pressure dependent contractions ( 261 ). Studies of human thoracic duct indicate many conserved physiological mechanisms between species including L-NAME- and indomethacin-sensitive regulation of contractile behavior ( 289 ) as well as pressure-dependent, pH-sensitive regulation of contractility ( 290 ). Isolated mouse lymphatic collecting vessels are now a significant portion of the current lymphatic isolated vessel zeitgeist. Contractile function has been assessed in mouse isolated lymphatic collecting vessels from a variety of anatomical regions, including afferent and efferent popliteal lymphatic vessels, axillary afferent lymphatic vessels, inguinal efferent lymphatic vessels, inguinal axillary lymphatic vessels, mesenteric lymphatic vessels, superficial and deep cervical lymphatic vessels, illiac efferent lymphatic vessels, and thoracic duct among others ( 261 ) ( 165 ) ( 274 ). However, there has not been systematic experimentation to test for flow-mediated inhibition in the majority of the lymphatic vessels from the aforementioned tissues. Testing lymphatic endothelial function in the mouse visceral cavities is complicated by the limited spontaneous tone development and lack of regular robust contractions. The mouse thoracic duct and mesenteric vessels are responsive to norepinephrine ( 261 ), although LMCs have store operated Ca 2+ entry, which can cause vasospasm and complicate preconstriction with IP 3 generating compounds such as U46619 or ET-1 ( 291 ). While instructions for the isolation for each of these vessels are readily available, it cannot be overstated how delicate these thin-walled vessels in mice are and the need for a delicate microdissection approach to retrieve healthy and functional vessels for isolated vessel myography. In no circumstances should the vessel be stretched away from the tissue and pulled out of solution as it will surely incur stretch-induced damage and disrupt the fine electrical LEC:LEC or LMC:LMC electrical coupling that is required for normal pressure-dependent contractility or flow-mediated and agonist stimulated inhibition experiments. Similarly, care must be made when connecting the pipettes to the fluid columns and adjusting the reservoir heights to prevent large pressure spikes which, along with prolonged supraphysiological pressures, can also damage the vessels. Rodent models expressing genetically encoded fluorescence reporters driven by expression of the known LEC markers, Prox1 and Vegfr3 ( 292 )( 293 )( 294 )( 295 )( 296 ) can be used to visualize and assist in identifying lymphatic vessels in any anatomical region of interest. Dissection Tools: The following list includes some of the dissection instruments that can be employed for isolation of lymphatic vessels: Ultra Fine Forceps for handling and pinning the perilymphatic tissue- 0.1 x 0.06 mm (For example see FST Cat.) Sharpened to 25-50 μm width 10 μm depth for tissue handling. Sharpened to 10-20 μm width <10 μm depth cleaning the vessel. Scissors for cutting the skin- 9 cm Cutting edge: 16 mm Spring Scissors for cutting the perilymphatic tissue out- 9 cm Cutting edge: 5 mm Tip Ø: 0.35 mm (for example see FST Cat. #91500-09). Vannas-Tübingen style or similar fine Spring Scissors for lymphatic vessel cleaning: - 8.5 cm Cutting edge: 5 mm Tip Ø: 0.075 mm (for example see FST Cat. #15003-08). Moria style or similar fine Spring Scissors for lymphatic vessel cleaning: 8 cm Cutting edge- 5 mm Tip Ø: 0.15 mm (for example see FST Cat. #15396-00). Pressure Myography: Functional and structural characterization of isolated lymphatic vessels requires fine intraluminal pressure control. Automated pressure control can be achieved by microfluidic systems, e.g., ElveFlow OB1; however, water columns can be employed for manual or automatic/semi-automatic pressure control. Due to the limited pressures under which the lymphatic collecting vessels and ducts normally operate, digital height gauges (Mitutoyo 570-414 HDS-H24"CX Digimatic HDS Height Gage, 24"/600 mm, 0.0005"/0.01 mm) can be adapted for fine control of the water column height (0.01 mm) and to avoid pressure spikes while making reservoir height changes. Prior to testing flow mediated or agonist stimulated inhibition of contractions, the vessel should be assessed for damage which could confound the data. Upon cannulation, the vessels should be able to inflate and tested for leaks in case a small transparent pre-collector or capillary branch was cut. Leaks can often be visualized by Schlieren lines, if BSA supplemented Krebs from the lumen is leaking into the albumin lacking perfusing buffer or when movement of small particles or cells is observed. Occasionally, lymphocytes still trapped inside the lumen can also help determine if leaks are present although lymph should be flushed to prevent unnecessary confounding actions of the lymphocytes. It is exceptionally critical that the micropipettes and all the tubing lines to the reservoir (especially any valves in the path) are cleared of bubbles as imposed flows can result in bubbles clogging the pipettes (especially at low pressures) and thus preventing flow or incidental damage or denudation if they pass into the vessel lumen. While variable across the vessels, sustained high lymphatic pressures in mice should be limited to 10-12 cmH 2 O (pressures at which contractions are usually insufficient to alter diameter) before irreversible damage is done, often indicated by an elevated vessel tone and multiple pacemaker activity when returned to lower pressure. Also, one should look for multiple contraction initiation sites and whether contraction waves are fully conducted along the length of the vessel, as these observations can explain potential confounders of both amplitude and frequency. Once the vessel is cannulated and connected to the fluid reservoirs, the regulation of pressure by altering the reservoir height should be tested again and briefly set to a pressure of 8 cmH 2 O to remove the slack from the vessel. Care must be made to not overstretch the vessels ( 268 )( 297 ) which will not only confound contraction frequency and amplitude but also alter the passive diameter and tone calculation. After 30-45 minutes of equilibration at 37°C at a moderate pressure 2-3 cmH 2 O, most lymphatic vessels will have a regular rhythm (noted by a consistent end diastolic diameter, i.e., EDD) and the contraction wave will typically start at one of the ends and propagate fully to the other often in a retrograde direction. Vessels with multiple pacemakers were often damaged during dissection or over stretched and will have colliding contraction waves which will result in abnormally low contraction amplitude and frequency depending on the tracking site. The damage induced pacemaker site may be refractory to the endothelial signaling and confound the experiment. Assessment of Endothelial-Dependent Regulation of Lymphatic Vessel Function: Testing lymphatic endothelium-dependent regulation of isolated lymphatic collecting vessels has been primarily accomplished through either flow-by imposed pressure gradients, by a concentration response curve to Ach, or by activation of endothelial TRPV4 channels ( 280 ). Similar to the arteriolar vasculature, the lymphatic endothelium can utilize multiple signaling modalities to regulate the contractile function of their ensheathing musculature such as NO ( 271 ), arachidonate metabolites ( 286 )( 298 )( 278 ), and histamine ( 299 ). Flow Mediated Inhibition of Contractions: As most healthy lymphatic vessels develop regular spontaneous contractions, there is little need for preconstriction, though the midpoint pressure should be set at a pressure (3-5 cmH2O) that induces high frequency contractions sufficient for evaluating the dynamic range of inhibition and imposed flows. It is imperative that micropipette flow resistance is matched to limit a confounding pressure change artifact. Ideally, the same pipettes can be used for the majority of the vessels included in the study barring significant diameter changes. For rat mesenteric and mouse popliteal lymphatic vessels, a tip diameter of 80-100 μm is used, while large lymphatic ducts like the thoracic duct will require larger pipettes of about 200-300 μm ( 260 ). For large diameter vessels with high flow rates the reservoir bath heights should be checked throughout the experiment. An inline heater or other mechanisms to warm the inflow PSS line should be considered as temperature can be a confounding variable on contraction frequency. Vessel tracking should be performed near the midpoint of the vessel where the pressure will be maintained but should avoid the valve sites if possible. Imposed flow should be set by a change in reservoir height of equivalent magnitude but opposite direction and in the direction of physiological flow due to the presence of the secondary valves which would close and prevent flow if the output was raised while the input was lowered. Flow changes can develop slowly, and the vessel should be tracked for at least 5 minutes at each imposed flow setting before returning to the equilibration condition. Flow responses can then be repeated in the presence of 100 μM L-NAME, 3 μM indomethacin, or 10 μM α-methyl-DL-histidine dihydrochloride. Finally, an NO donor such as 100 μM sodium nitroprusside should be applied at the end of the experiment to ensure LMC reactivity to NO signaling was intact. While analyzing the contraction frequency, contraction amplitude, and vessel tone, it is worth noting that in some lymphatic vessels (such as the mouse inguinal axillary vessel), the action potential cycles are faster than the physical kinetics of the contraction and relaxation cycle. This results in overlapping contractions and a non-stable “EDD” which is now a function of the electrical pacemaking as opposed to steady state tone. Thus, an elongation of the electrical cycle (i.e. reduction in contraction frequency) invariably results in a higher EDD before the next contraction starts and would be calculated as an apparent reduction in developed tone, however this should be considered thoughtfully in the analysis. Passive diameters can be generated at each of the differential pressures to appropriately account for a tracking site offset from the midpoint. Acetylcholine Stimulated Inhibition of Contractions: Lymphatic endothelial function can also be assessed from the concentration response relationship to ACh in vessels mounted under isobaric conditions. In mice, ACh induced inhibition of contraction amplitude and frequency appears to be completely dependent on NO production ( 300 )( 301 ). An ACh concentration response from 1 nM to 1 μM will provide the full dilatory range and at the higher concentrations explore excitatory stimulation of muscarinic receptors on the muscle cells. ACh should be premixed into 60-100 μL of buffer to help mitigate acute high concentration exposure while mixing. ACh should be added to the bath in parallel or near parallel to the vessel to prevent damage from overstretching the vessel when mixing. Pipetting should be controlled to limit the generation of bubbles which can stick to the vessel or float into the light path and obfuscate the tracking. As with the flow induced inhibition, a 20-minute equilibration with 100 μM L-NAME followed by a second ACh response should be performed with L-NAME supplemented perfusion. Passive diameters can then be recorded and changes in contraction frequency and tone calculated. Quantitative Assessment of Lymphatic Valve Function: Lymphatic valves are bicuspid structures where each leaflet is composed of a specialized extracellular matrix scaffold (i.e., collagen and elastin) that is surrounded by monolayers of LECs ( 302 ). Competent valves are critical for the efficient transport of lymph, as these structures not only minimize retrograde lymph flow, but also allow for the generation of propulsive pressure during a contraction cycle. This is particularly important in anatomical regions where lymph transport must be accomplished against an adverse pressure gradient, e.g., as that imposed by gravitational forces. Lymphatic valve structural abnormalities may be identified and characterized by immunofluorescence confocal imaging of isolated, cannulated, and pressure-fixed lymphatic vessels ( 303 )( 304 ); however, these approaches do not provide insights into the functional status and competency of the valves. Lymphatic valve function can be assessed in isolated and cannulated lymphatic vessels by two tests: valve gating and backleak tests ( 305 )( 306 ). Assessment of Lymphatic Barrier Function Once interstitial fluid enters the lymphatic networks, lymph is meant to stay within the lymphatic networks. Lymphatic permeability in collecting lymphatic vessels is mainly regulated by endothelial junction integrity. The isolated lymphatic vessel preparation provides an ideal experimental approach for quantitative assessment of lymphatic permeability, one that incorporates not only the main, endothelial component, but also the remaining constituents of the lymphatic wall ( 307 )( 308 ). Lymphatic permeability is calculated by determining the efflux of BSA, or other solutes, across the lymphatic wall at a set level of transmural pressure. As an example, a controlled BSA gradient can be established by using a physiological buffer containing 1% BSA inside the vessel lumen (where 1/20-th of the BSA is conjugated with a fluorescent dye, e.g., Alexa-488), while the vessel is immersed in a bath containing a physiological buffer with 0.1% non-fluorescent BSA. Although fluorescent BSA conjugates are commercially available, molecular size characterization of the employed BSA is fundamental for accurate assessment of lymphatic permeability, this can be achieved by generating Alexa-488 conjugated BSA in house. Briefly, Alexa-488 and BSA are allowed to react at room temperature for 30 minutes, followed by filter centrifugation to limit maximum molecular size (e.g., 30 kDa), and subsequent removal of free dye by buffer exchange column chromatography. An in-depth description and discussion of this technique has been recently provided by the original authors ( 301 )( 309 ). Characterization of Intracellular Ca 2+ Activity in the Lymphatic Endothelium: Assessment of intracellular Ca 2+ changes in the lymphatic endothelium provides valuable insights into the mechanistic processes that regulate the different aspects of collecting lymphatic function. Please refer to the earlier section “ EC Ca 2+ Imaging ” for information on imaging of intracellular Ca 2+ in vascular structures, including lymphatics. J. Studying Ion Channels in Native ECs using patch-clamp electrophysiology ECs are continuously exposed to mechanical forces, such as shear stress from blood flow and cyclic stretch, as well as chemical stimuli, including blood-borne factors and signaling molecules ( 310 )( 311 ). Ion channels are critical transducers of these stimuli, modulating endothelial membrane potential and intracellular Ca 2+ signaling to mediate physiological responses. ECs express a diverse array of potassium (K + ), chloride (Cl − ), and transient receptor potential (TRP) cation channels ( 312 )( 313 ) ( 314 ) Dysregulation of endothelial ion channel activity has been implicated in the pathogenesis of cardiovascular and cerebrovascular diseases. For example, abnormal channel function impairs endothelial-dependent vasodilation ( 315 ), contributing to hypertension and atherosclerosis ( 316 )( 317 )( 318 ). In the cerebral vasculature, ion channel dysfunction has been associated with neurovascular disorders, including Alzheimer’s disease and cerebral small vessel disease ( 319 )( 320 ). Given their critical role in endothelial physiology and vascular homeostasis, endothelial ion channels have emerged as potential therapeutic targets for vascular diseases. Therefore, studying these proteins using methodologies that closely mimic physiological conditions will provide deeper insight into their role in vascular function and disease mechanisms. Consequently, the electrophysiological investigation of endothelial ion channels has become an essential and rapidly expanding field, offering critical insights into the molecular underpinnings of endothelial dysfunction and identifying new avenues for targeted therapeutic intervention. Patch-clamp electrophysiology remains the “gold standard” for studying ion channels, enabling precise characterization of single-channel kinetics, biophysical and pharmacological properties, as well as physiological and pathological regulation. Culturing ECs, even primary cells, induces significant transcriptomic alterations ( 124 ) , which can confound the interpretation of ion channel function. Therefore, utilizing freshly isolated native ECs is essential for obtaining rigorous data. Combining this electrophysiological technique ( 321 ) with protocols for isolating native (uncultured) ECs, patch-clamp electrophysiology offers an invaluable approach for studying ion channel function in a physiologically relevant context. In the following sections, we will describe the essential components of electrophysiological rigs, the different configurations of the patch-clamp technique, and key experimental considerations to optimize recordings from native ECs. Electrophysiological Rigs for Patch-Clamp: An electrophysiology rig is a specialized setup with several key components for controlling and measuring electrical signals in cells or tissue. Depending on the investigators' needs, additional elements may also be present. In this section, we will review the essential components ( Fig. 14 ). Figure 14. Open in a new tab Components of an electrophysiological rig. (A) Antivibration table. (B) Faraday cage. (C) Horizontal flaming/brown micropipette puller. (D) Microforge. (E) Electrode with the pipette holder. (F) Headstage. (G) Main amplifier. Critical switches are highlighted in dotted red squares, such as Pipette offset, “Zap” button for whole-cell facilitation, Pipette capacitance compensation, Whole-cell parameters, and Mode controller. (H) Microscope. (I) DC power supply. (J) Micromanipulator. a. Minimizing Environmental Noise Effective isolation from environmental vibrations and electrical or mechanical noise is essential for obtaining high-quality electrophysiological recordings. An anti-vibration table is a critical component that isolates the experimental setup from mechanical disturbances ( Fig. 14A ). In addition to mechanical stability, electromagnetic noise must be eliminated to prevent interference with electrophysiological measurements. This is achieved using a Faraday cage ( Fig. 14B ), a wire-mesh enclosure that surrounds the sensitive components of the experimental setup, including the microscope and electrode. b. Microelectrode Fabrication Patch-clamp electrophysiology requires the fabrication of glass micropipettes with a 1-3 μm tip diameter. The choice of pipette material depends on the experimental requirements, with borosilicate glass tubing being the most used. These pipettes typically have a wall thickness of 0.2–0.3 mm, an outer diameter (O.D.) of 1.5 mm, and an inner diameter (I.D.) of 0.86 mm or 1.10 mm. Glass pipettes typically contain an internal filament to facilitate the loading of the internal solution to the fine tip by capillary action. Micropipettes are fabricated using a micropipette puller, which heats and pulls glass tubing to create the desired tip shape ( Fig. 14C ). A microforge is used to reshape and polish pipette tips, ensuring optimal recording properties ( Fig. 14D ). The microelectrode holder ( Fig. 14E ) connects the microelectrode to the headstage ( Fig. 14F ). After the microelectrode is filled with internal solution, it is mounted on the holder, which contains a silver/silver chloride (Ag/AgCl) wire. This wire facilitates a reversible redox reaction, where AgCl dissociates into silver (Ag + ) and Cl − ions, maintaining electrical stability. For accurate recordings, the potential difference between the recording electrode and the ground electrode (connected to the headstage) should be zero when both are immersed in electrically connected solutions with equal chloride concentrations. AgCl depletion can occur due to prolonged exposure to high electrolyte concentrations or excessive operational voltages, leading to electrode polarization and unstable recordings. A color change from dark gray to silver indicates AgCl depletion. To restore function, “rejuvenate” the wire electrode by immersing it in sodium hypochlorite (bleach) for 10-20 minutes until the Ag/AgCl coating is fully restored. c. Amplifier The headstage is connected to a patch-clamp amplifier ( Fig. 14G ), enabling precise measurement and control of ionic currents across biological membranes. The amplifier magnifies picoampere (pA) currents recorded from cells while minimizing electrical noise. In voltage-clamp mode, the amplifier injects current to maintain a stable membrane potential. To improve recording fidelity, modern patch-clamp amplifiers incorporate advanced compensation mechanisms that correct for distortions caused by the electrical properties of the recording system ( Fig. 14G ). d. Software Commercially available software allows the visualization and analysis of the data recorded. For example, Clampex (Molecular Devices) provides control over data acquisition. Clampex works seamlessly with Clampfit (Molecular Devices), which is designed for offline analysis. PatchMaster (HEKA Instruments) serves a similar function. e. Microscopes Inverted microscopes are typically used for patch-clamp electrophysiology in isolated cells ( Fig. 14H ), whereas upright microscopes are used for recording from brain slices and other intact tissues. A 60X objective (or higher) is useful for visualizing ECs on an inverted microscope. Some illumination sources commonly integrated into microscopes for patch-clamp include LED-based systems such as the CoolLED pE-Series (CoolLED. Andover, UK). Alternating current (AC)-powered illumination sources introduce 50 (Europe)/60 (USA) Hz cycle electromagnetic noise, which can be eliminated by using a direct current (DC) power supply to power the illumination system ( Fig. 14I ). A micromanipulator precisely controls the three-dimensional movement of the headstage and the pipette ( Fig. 14J ). h. Perfusion and Temperature Control Systems for Patch-Clamp Electrophysiology A perfusion system is essential for delivering extracellular solutions to the recording chamber during patch-clamp experiments. These systems are typically constructed using commercially available multi-channel manifolds and Teflon tubing, which minimizes the absorption of drugs and other chemicals. Fluid flow is regulated by manual valves or electronically controlled solenoids, enabling rapid and precise switching between solutions. Gravity-driven perfusion is the simplest and most cost-effective approach, while pressure-driven and peristaltic pump systems offer faster and more consistent solution exchange. Specific applications require precise and rapid solution changes. This can be accomplished using specially built rapid stepper instruments (i.e., SF-77B, Warner Instruments) or picospritzer systems coupled with fine glass capillaries, allowing fast and spatially confined agonist delivery. Patch-clamp experiments are routinely conducted at room temperature. However, for some studies, maintaining physiological temperature during recordings is critical ( 322 ). Patch-clamp configurations: The patch-clamp technique encompasses several configurations, cell-attached, inside-out, whole-cell, perforated whole-cell, and outside-out, each offering distinct advantages for studying ion channel function, regulation, and pharmacology under physiological and controlled experimental conditions. A brief overview of these configurations is provided below. a. Cell-attached Patch-clamp The cell-attached patch-clamp configuration is widely used to investigate ion channel activity while maintaining the integrity of the intracellular environment. In this configuration, a glass micropipette containing an ionic solution is positioned in contact with the cell membrane. Mild negative pressure is applied to establish a gigaseal between the pipette and the membrane ( Fig. 15A ), isolating a membrane patch within the pipette tip. Currents from the ion channels in this region can be recorded. Because the intracellular environment remains unperturbed, this technique is advantageous for examining ion channel activity under physiological conditions and in response to mechanical stimuli applied to the plasma membrane. Figure 15. Open in a new tab Patch-clamp configurations. (A) Cell-attached configuration is achieved by applying mild suction to the plasma membrane, getting a gigaseal. (B) If the pipette is retracted during a cell-attached configuration, the membrane patch will also be removed from the cell, resulting in the inside-out configuration. (C) If strong suction is applied during cell-attached mode, the plasma membrane will break, allowing access to the cytoplasm, a configuration known as whole-cell. (D) The perforated patch configuration is obtained by adding an ionophore into the pipette solution on a cell-attached mode. (E) Outside-out configuration can be achieved by pipette retraction during a whole-cell mode. b. Inside-Out Patch-Clamp The inside-out patch-clamp configuration is achieved by forming a gigaseal in cell-attached mode, followed by rapid withdrawal of the pipette, which detaches a small patch of membrane from the cell and exposes its intracellular surface to the bath solution ( Fig. 15B ). This allows precise control over the composition of the intracellular environment, making it particularly useful for investigating the effects of Ca 2+ and other second messengers on ion channel function. c. Conventional Whole-Cell Patch-clamp The conventional whole-cell patch-clamp configuration is achieved by forming a gigaseal between the pipette and the cell membrane, followed by applying sufficient suction to rupture the membrane patch within the pipette tip, dialyzing the cell with the internal solution ( Fig. 15C ). This configuration enables precise control of the intracellular environment, including ionic conditions, while recording whole-cell currents ( 323 )( 324 ). However, whole-cell configuration is not ideal for studying intracellular regulation by second messengers, as the pipette solution replaces the native cytoplasmic content. d. Perforated Whole-Cell Patch-Clamp A variation of this configuration is the perforated patch, in which a pore-forming (ionophore) compound is added to the pipette solution ( Fig. 15D ). This results in the perforation of the membrane, enabling voltage clamp while preserving cytoplasmic content. Pore-forming molecules are selective for specific ions. For example, amphotericin B and nystatin allow monovalent ions to permeate while remaining impermeable to divalent ions, thus preserving intracellular Ca 2+ signaling ( 325 )( 326 ) ( 327 ). Other ionophores, such as gramicidin, facilitate the movement of monovalent cations while excluding anions, making it appropriate for keeping intracellular Cl − concentration unchanged ( 328 ). The prolonged perforation time can be a limitation of this technique, as the time needed can vary between cell types and ionophore concentrations (ranging from 5 to 30 minutes). e. Outside-Out Patch-Clamp The outside-out patch-clamp configuration is achieved by first establishing a whole-cell configuration, followed by carefully retracing the pipette. This causes the membrane to reseal with the extracellular surface facing outward ( Fig. 15E ). This configuration allows precise control over the external solution, making it ideal for studying the effects of neurotransmitters, drugs, and extracellular ions on ion channel activity. Voltage vs. Current-Clamp: In the voltage-clamp configuration, the membrane potential is held at a predetermined value (the command potential) by injecting current while measuring ionic current. The command potential can be held at a constant level or applied at various levels as steps or a ramp. Current is plotted as a function of voltage to identify rectification properties and voltage-dependent activation or inhibition. Current-clamp measures the changes in membrane potential resulting from ion channel activity. This mode is commonly used for studying action potentials or excitability in excitable cells but can be used to measure the resting membrane potential of ECs ( 315 )( 329 ). Strategies for Isolating Specific Ion Channel Currents in Native ECs: Careful experimental design is essential for isolating specific ionic currents in native EC. Prior studies using heterologous expression systems provide key information on ion selectivity, voltage dependence, kinetics, and pharmacological modulators, serving as a foundation for optimizing recording conditions. Ion substitution is frequently employed to eliminate unwanted conductances. For example, cesium (Cs + ) can be substituted for K + to decrease currents through K + -selective channels ( 330 ), while large, non-permeant anions such as gluconate or aspartate suppress Cl − currents ( 331 ). Pharmacological inhibitors can further isolate currents. Tetraethylammonium (TEA) selectively blocks voltage-gated and Ca 2+ -activated K + channels ( 330 )( 332 ), whereas inwardly rectifying K + (K IR ) channels can be isolated using the selective blocker Ba 2+ ( 333 ). Intracellular Ca 2+ buffering is critical when studying Ca 2+ -sensitive ion channels using the conventional whole-cell configuration. Free Ca 2+ levels can be clamped at specific levels using selective Ca 2+ chelators such as EGTA (ethyleneglycol- bis(β-aminoethyl)-N,N,N',N'-tetraacetic acid) or BAPTA (1,2-bis(o-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid). Maxchelator (Stanford, USA) and PyChelator are open-source software tools that calculate intracellular free Ca 2+ levels ( 334 ). When designing external or internal solutions for electrophysiology, it is essential to maintain physiological osmolarity (280-310 mOsm/L), as this contributes to obtaining high-quality gigaseal. Theoretical osmolarity should first be calculated, followed by direct measurement using an osmometer. If the solution is hypoosmotic, an inert molecule such as mannitol or sucrose can be added to adjust osmolarity without altering ionic composition. In cases requiring ionic substitution, maintaining osmolarity and isotonicity is best achieved by reducing the concentration of the most abundant ions. For example, when increasing external K + , a proportional reduction in Na + helps preserve osmotic balance. Native EC isolation: ECs undergo rapid phenotypic changes under culture conditions ( 124 ), necessitating their use immediately after isolation for optimal experimental rigor. Cells obtained using this protocol remain viable for at least six hours. a. Isolation of Mesenteric Resistance Arteries A detailed protocol for the isolation of mesenteric arteries has been previously described ( 1 ). A critical step to be considered is the discrimination between veins and arteries, which can be observed by the large lumen size and thinner walls of veins compared to arteries ( 335 ). To ensure a sufficient number of viable ECs, isolating at least 6-8 third-order arteries is recommended ( Fig. 16A ). Please note that, in rodents (mice and rats), 3rd-order (and 4th-order) mesenteric arteries are widely accepted as resistance arteries, as their lumen diameter is typically <250 μm and they contribute substantially to peripheral vascular resistance. Transfer the isolated arteries to a centrifuge tube containing ice-cold EC isolation solution ( Table 8 ). The EC isolation solution is formulated specifically to facilitate the separation of ECs from underlying VSMCs while maintaining EC viability. Low Ca 2+ (0.1 mM) reduces Ca 2+ -dependent EC-EC junctional adhesion and minimizes Ca 2+ -dependent VSMC contraction, both of which aid in selective EC detachment. Higher Mg 2+ (2 mM) supports membrane stability during enzymatic digestion. Na-glutamate substitutes for part of the NaCl to maintain osmolarity while reducing extracellular chloride concentration. Lowering Cl − helps minimize VSMC depolarization and contractile activation during the isolation process, thereby facilitating gentler detachment of ECs. Figure 16. Open in a new tab Isolation of Endothelial Cells from Different Vascular Beds. (A) Arteries are isolated from mouse mesenteries. After careful dissection and cleaning, the arteries undergo enzymatic digestion and mechanical dispersion to obtain isolated mesenteric endothelial cells. A representative mesenteric endothelial cell is shown on the right (scale bar: 10 μm). (B) Cerebral pial arteries are collected from the brain. Following enzymatic digestion and mechanical dispersion, isolated cerebral artery endothelial cells are obtained. A representative cerebral artery endothelial cell is shown on the right (scale bar: 10 μm). (C) To isolate brain capillary endothelial cells, surface arteries from the brain are removed, and three 1-mm thick cortical slices (dotted lines) are excised and homogenized. Capillaries are collected by filtration and subjected to enzymatic digestion and mechanical dispersion to obtain individual brain capillary endothelial cells. A representative brain capillary endothelial cell is illustrated on the right (scale bar: 10 μm). All images were captured using differential interference contrast (DIC) microscopy at 40X magnification. Created in BioRender. Lavanderos, B. (2026) https://BioRender.com/43ip1tb . Table 8. Concentrations of compounds in EC Isolation Solution used during isolation process. EC Isolation Solution (pH 7.3 adjusted with NaOH) Concentration (mM) NaCl 55 mM KCl 6 mM MgCl 2 2 mM CaCl 2 0.1 mM Glucose 4 mM HEPES 10 mM Na-glutamate 80 mM Open in a new tab b. Isolation of Cerebral Pial Arteries Cerebral pial arteries, the anterior, middle, and posterior cerebral arteries, originate from the circle of Willis and branch into smaller vessels along the surface of the brain. A standardized protocol for pial artery isolation was described in our previous guidelines ( 1 ). All the cerebral pial arteries from a brain should be collected for the isolation of ECs to ensure that a sufficient number of cells are obtained. ( Fig. 16B ). The dissected vessels should be transferred into a centrifuge tube using a capillary glass pipette, and the solution should be replaced with ice-cold EC isolation solution ( Table 8 ). c. Isolation of Brain Capillaries To isolate brain capillaries, first remove large surface vessels from the brain using a moistened cotton swab. Place the brain in a brain matrix slicer (Fisher Scientific, Hampton, NH, USA) and cut 1-mm thick coronal sections. Collect three slices from the cortical region ( Fig. 16C ) and immediately transfer them to a Dounce homogenizer containing 10-15 mL of artificial cerebrospinal fluid (aCSF) ( Table 9 ). The aCSF should be pre-aerated for 20–30 minutes with a gas containing 21% O 2 , 5% CO 2 , balanced with N 2 . Table 9. Composition of artificial cerebrospinal fluid (aCSF) used for brain homogenization during brain capillary EC isolation. aCSF Concentration (mM) NaCl 124 mM KCl 3 mM MgCl 2 2 mM CaCl 2 2 mM Glucose 4 mM NaHCO 3 26 mM NaH 2 PO 4 1.25 mM Open in a new tab Gently homogenize the tissue using a Dounce homogenizer until no visible tissue fragments remain, minimizing bubble formation to prevent damage to the capillaries. To isolate capillaries, slowly filter the homogenate dropwise through a 70-μm mesh filter ( Fig. 16C ), ensuring even distribution by moving the pipette across the filter surface to prevent clogging. To collect the capillaries, invert the filter over a fresh 50-mL tube and release the captured networks by flushing with EC isolation solution ( Table 8 ) using vigorous pipetting. d. EC dispersal Individual ECs are prepared from isolated arteries or capillaries by enzymatic digestion followed by mechanical dispersion. Initially, vessel segments are digested using 0.5 mg/mL neutral protease and 0.5 mg/mL elastase (Worthington Biochemical Corporation, Lakewood, NJ, USA) in EC isolation solution at 37°C for 15 minutes. Following digestion, the arteries are washed with ice-cold EC isolation solution and then incubated for 2 minutes at 37°C with 0.5 mg/mL collagenase type I (Worthington Biochemical Corporation) ( 104 )( 315 )( 336 ). The tissue is gently centrifuged at 420 g for 5 minutes at 4°C and triturated using a polished glass Pasteur pipette to release individual ECs ( 323 )( 336 ). After mechanical dissociation of mesenteric and pial arteries, some residual connective tissue may remain, whereas brain capillary EC suspensions should be free of connective tissue contamination. EC morphology varies across vascular beds ( Fig. 16 ), with cerebral pial and mesenteric ECs exhibiting capacitance values of approximately 12 and 9 picofarads (pF), respectively. In contrast, brain capillary ECs are smaller, with an average capacitance of 6-8 pF. Enzymatically dispersed artery and arteriole suspensions contain a mixture of VSMCs and ECs. Intact VSMCs are elongated when relaxed, making them easily distinguishable from ECs. However, damaged or contracted VSMCs may resemble ECs, complicating identification. To improve cell-type discrimination, transgenic endothelial reporter mice ( 315 ) or EC-specific dyes such as Isolectin B4 (ThermoFisher, Waltham, MA, USA) can be used. Additionally, electrophysiological characterization can provide functional distinction. For example, ECs express large-conductance Ca 2+ -activated potassium (BK) channels at much lower levels than VSMC ( 337 ). Example Experiments: To illustrate the above techniques, we provide two example protocols. Example 1: Whole-cell K IR 2.1 currents in native brain capillary ECs. This section provides a step-by-step protocol for establishing the whole-cell patch-clamp configuration, using K IR 2.1 currents from native brain capillary ECs as an example. 1. Cell Preparation and Chamber Setup Add an aliquot of freshly isolated ECs to the recording chamber and allow them to attach for 15 minutes. To remove debris and non-adherent cells, perfuse the chamber with bath solution for 2–5 minutes. Adherent brain capillary ECs are elongated, with the nucleus slightly off-center ( Fig. 16C ). 2. Pipette Preparation and Giga-Seal Formation Fill the patch pipette with the internal solution ( Table 10 ) using a Microfil needle (28-gauge, 67 mm). To remove air bubbles, gently tap the pipette and insert it into the pipette holder ( 338 ). Lower the pipette and, using the amplifier’s pipette offset function, adjust the baseline to 0 pA ( Fig. 14G ). When asymmetric ionic solutions are used, like in this case, a liquid junction potential (LJP) is generated at the interface between pipette and bath solutions due to differences in ionic mobility. This potential is not eliminated by the pipette offset function and should be accounted for when interpreting absolute membrane voltages. Investigators should calculate the LJP using tools such as LJPcalc, and then either compensate for it electronically (online correction) or subtract the calculated value during analysis (offline correction) ( 339 ) ( 321 ). In this case, using an organic anion-based internal solution, the calculated LJP is −13.7 mV. Lower the pipette and lightly touch the cell membrane, then apply a slight negative pressure (e.g., by mouth or with a syringe) to bring the membrane into contact with the pipette tip. As the high-resistance seal forms, the resistance will gradually increase from megohms (MΩ) to gigohms (GΩ). The decay of capacitive transients should return to baseline levels ( Fig. 17A ). Cells with leaky seals will have excessive baseline current and should be discarded. Use the pipette capacitance compensation function on the Axopatch amplifier ( Fig. 14G ) to minimize capacitive transients during the membrane test ( Fig. 17A ). Table 10. Composition of internal solution used for the recording of K IR channel currents in whole-cell configuration. Pipette solution (pH 7.2 adjusted with KOH) Concentration (mM) K-aspartate 110 mM NaCl 10 mM KCl 30 mM MgCl 2 1 mM HEPES 10 mM Open in a new tab Figure 17. Open in a new tab Achieving Whole-Cell Access. (A) Representative example of transient capacitive currents and resistance changes during gigaseal formation. Upon contacting the cell, pipette resistance (Rt) increases, and applying gentle suction facilitates membrane attachment, leading to a seal with gigohms (GΩ) resistance. The formation of a high-resistance seal generates transient capacitive currents. (B) Once the whole-cell configuration is established, membrane capacitance (Cm) increases, and access resistance (Ra) decreases. Adjusting the whole-cell capacitance compensation command in the amplifier reduces transient capacitive currents, optimizing signal resolution. However, overcompensation may introduce artifacts, causing deflections in residual capacitive currents, potentially leading to underestimation or distortion of recorded events. 3. Membrane Rupture and Whole-Cell Access Once a high-resistance seal is achieved, gently apply suction to rupture the membrane and establish whole-cell configuration. If mechanical rupture affects cell viability, use the amplifiers “Zap” function ( Fig. 14G ) to apply a brief electrical pulse to break the membrane. Upon successful rupture, membrane capacitance (Cm) should increase from ~12 pF to ~20 pF ( Fig. 14B ). Allow 3–5 minutes for the pipette solution to dialyze the cell before beginning recordings. Use the whole-cell compensation settings on the amplifier to adjust for membrane capacitance, avoiding overcompensation, which can lead to underestimation of ionic currents ( Fig. 17B ). Achieving acceptable access resistance depends on a high-quality gigaseal, as well as cell viability, the relative osmolarity of internal and external solutions, and mechanical stability. 4. Electrophysiological Recording K IR 2.1 channels are activated by extracellular K + and strongly inhibited by micromolar concentrations of Ba 2+ . A voltage ramp protocol is commonly used to record K IR 2.1 currents. For this example, the membrane potential was held at −50 mV between sweeps and then ramped from −100 mV to +40 mV over 400 ms ( Fig. 18A ). Ramps were repeated every 2 seconds throughout the recording. Currents are initially recorded from cells bathed in a solution with physiological external [K + ] (6 mM, Table 11 ) ( Fig. 18B - C ). K IR 2.1 channels are inactive, and currents are essentially zero pA at all command potentials ( Fig. 18B - C ). After approximately 30 s, the chamber is perfused with a high K + (60 mM, Table 12 ) bathing solution to activate K IR 2.1 channels. Approximately one minute is required for solution exchange and currents to reach a steady state ( Fig. 18B - C ). Under these conditions, inward current amplitude increases ~8-fold compared to low K + conditions at −100 mV. After the K IR 2.1 current reaches its maximum in high K + , a Ba 2+ (10 μM) solution is perfused to fully inhibit the current. This solution induces complete current inhibition of the inward current within 1 minute ( Fig. 18B - D ). To analyze K IR 2.1 currents, individual current sweeps are selected, as shown in Fig. 18C - D . The Ba 2+ -sensitive current, representing the total KIR2.1 current, is obtained by subtracting the steady-state current following Ba 2+ application from the maximal current recorded in a high-K + bathing solution. For accurate comparisons between experimental conditions or groups, normalization to cell size is essential. This is achieved by dividing the recorded current by the cell capacitance, yielding current density, expressed as pA/pF. In wild-type, young adult mice, it have been reported current densities ranging from −15 to −8 pA/pF at −100 mV ( 320 )( 323 ). Figure 18. Open in a new tab Example whole-cell patch-clamp recordings of K IR 2.1 currents in native brain capillary endothelial cells. (A) Voltage ramp protocol used to measure whole-cell K IR 2.1 currents. (B) Representative time course of the activation and inhibition of K IR 2.1 current density in brain capillary endothelial cells at −100 mV. Thirty seconds of basal levels were recorded under physiological external K + concentrations (6 mM). Next, K IR 2.1 currents were activated by increasing external K + concentrations (60 mM). After reaching the maximal current, Ba 2+ (10 μM) was perfused to inhibit the K IR 2.1 current. Arrowheads indicate the points used in (C) and (D), where current density was plotted as a function of voltage. The color code represents individual traces for the indicated time. The difference between traces shown in (D) reflects the Ba 2+ -sensitive current. Table 11. Composition of physiological [K + ] bathing solution used for the recording of K IR channel currents in whole-cell configuration. Physiological [K + ] solution (pH 7.4 adjusted with NaOH) Concentration (mM) NaCl 134 mM KCl 6 mM CaCl 2 2 mM MgCl 2 1 mM Glucose 10 mM HEPES 10 mM Open in a new tab Table 12. Composition of high [K + ] bathing solution used for the recording of K IR channel currents in whole-cell configuration. High [K + ] solution (pH 7.4 adjusted with NaOH) Concentration (mM) NaCl 80 mM KCl 60 mM CaCl 2 2 mM MgCl 2 1 mM Glucose 10 mM HEPES 10 mM Open in a new tab Example 2: Measuring EC resting membrane potential using the current-clamp configuration. EC resting membrane potential can be measured using the current-clamp configuration. Most studies report that the membrane potential of ECs is between −20 and −40 mV ( 315 )( 340 ). The resting membrane potential of ECs can be measured using the current-clamp configuration as previously described ( 315 ). Before starting the experiment, configure the data acquisition system for current-clamp mode. Use a gap-free acquisition protocol with zero current injection to ensure continuous data recording. The cell-attached configuration is used to record membrane potential in current-clamp mode. Add an aliquot of freshly isolated ECs to the recording chamber and allow them to attach for 15 minutes. Wash the chamber by perfusing with the bathing solution to remove debris and non-adherent cells. A bath solution can be used to measure resting membrane potential ( Table 11 ). Please note that, while some solution is not fully physiological, such compositions are sometimes necessary to optimize stability, seal quality, or ion current isolation in patch-clamp recordings. Fill the patch pipette with internal solution ( Table 13 ) using a Microfil needle (28-gauge, 67 mm). To eliminate air bubbles, gently tap the pipette before inserting it into the pipette holder. Lower the pipette until it contacts the cell membrane, then use the Pipette Offset function on the amplifier ( Fig. 14G ) to adjust the baseline to 0 pA. Pipette resistance gradually increases upon membrane contact ( Fig. 17A ). Apply slightly negative pressure using suction or a syringe to facilitate tight pipette attachment to the plasma membrane. As the gigaseal forms, resistance should progressively increase from MΩ to GΩ. Capacitive transients will become evident in the membrane test screen of Clampex ( Fig. 17A ). Once the gigaseal stabilizes (>2 GΩ) for at least 3–5 minutes, switch the amplifier mode from Voltage-Clamp (V-CLAMP) to Current-Clamp (I-CLAMP NORMAL) on the Axopatch amplifier ( Fig. 14G ). Capacitive current compensation is not required in current-clamp mode, as the amplifier’s automatic series resistance compensation function ensures optimal recording conditions. Table 13. Concentrations of components of internal solution for measure of resting membrane potential in cell-attached configuration. Pipette solution (pH 7.2 adjusted with KOH) Concentration (mM) NaCl 10 mM K-aspartate 110 mM MgCl 2 1 mM CaCl 2 0.5 mM HEPES 10 mM K 2 HPO 4 4 mM EGTA 5 mM Open in a new tab K. EC Stiffness - Atomic Force Microscopy A more recent area of interest is related to the impact of mechanics and EC stiffness on vascular cell function, including EC function, under normal and diseased conditions. The most direct and sensitive method by which to assess these is via the use of atomic force microscopy (AFM), which allows one to elucidate cellular topographical changes and underlying cellular and tissue-level stiffnesses. For example, early AFM applications on ECs demonstrated that shear stress induces reorganization of the surface topography in ECs aligned in the direction of flow ( 341 ) and that EC cytoskeleton and stiffness can be modulated by substrate stiffness ( 342 ). In addition to assessing cellular stiffness, AFM has increasingly been applied to directly interrogate the endothelial glycocalyx. Through nano-indentation and polymer-brush modeling approaches, AFM can quantify glycocalyx mechanical parameters, including stiffness, brush “length,” and coverage, as well as topographical changes in response to physiological or pathological stimuli. Studies using cultured ECs and ex vivo vessels have shown that disease states such as diabetes and sepsis are associated with glycocalyx degradation and altered nanoscale mechanics, highlighting the utility of AFM for mechanobiological investigations of this critical endothelial structure ( 343 ) ( 344 ). AFM has also been used to interrogate vascular cell integrin signaling and mechanics. Indeed, integrins on ECs can be activated by both shear and integrin-mediated mechanisms to stimulate endothelin and other vasoactive agents in a myriad of vascular beds ( 345 ). These are just a few examples of the importance of EC mechanics and how AFM can help elucidate novel mechanical mechanisms of ECs in normal and diseased conditions. AFM Acquisition in EC Culture: While EC stiffness can be assessed in en face vessel preparations ( 346 ) and potentially in fixed tissue sections to assess relative differences in stiffness (not absolute), arguably the most reliable way to assess EC mechanics is by using isolated ECs in cell culture. AFM in live cell culture avoids the variability in tissue topography seen in en face preparations and the increases in absolute stiffness associated with fixed preparations. In a typical experimental protocol, plated live cells are placed onto an inverted microscope with a modified AFM stage, and an AFM head with an appropriate AFM cantilever is secured and placed on top of the dish of cells in order to conduct a nanoindentation protocol to assess EC stiffness. To reliably and rigorously assess EC and other vascular cell mechanics in live cell culture, there are several considerations below to ensure reliable and robust results: a. Tissue Culture Dish Size and Coating Considerations Prior to beginning, ensure that the size of the tissue culture dish onto which cells are plated can accommodate the AFM head/cantilever setup for your particular AFM. For example, the Asylum MFP-3D-BIO AFM cantilever holder can accommodate a 60 mm dish or larger. If your experimental setup requires smaller numbers of cells or if adequate cell availability is a consideration, ECs can be plated onto chamber slides, which can then be tacked inside a 100 mm tissue culture dish with media in order to perform AFM on cells. One thing to consider, depending on the scientific question being asked, is whether substrate stiffness matters for your experimental setup. Tissue culture plastic and glass are exceptionally stiff, but they are the most commonly used. One may also consider coating the plastic or glass with extracellular matrix proteins such as collagen, or something more inert such as gelatin to help cell adherence. Finally, there is ample data in the literature that the substrate stiffness upon which ECs are grown can also impact their inherent cellular stiffness ( 342 ). One may consider plating ECs onto polyacrylamide gels titrated to specific stiffnesses, depending on their experimental paradigm (commercially available from Matrigen, Irvine, California). b. Temperature AFM in EC culture is typically performed at room temperature ( 346 )( 347 ). While not ideal per se , it is a practical necessity. Even though some AFM systems are equipped with stage heaters to maintain temperature at physiological levels (~37°C), heating media/cells with the lid off for AFM access results in media evaporation and hypertonic solutions, quickly killing adhered cells/tissues. c. Cell Confluence The state of cellular confluence in the dish can influence cell mechanical properties ( 348 ). Typically, it is recommended that cells should be sub-confluent (50-80% range) to avoid contact inhibition, and AFM measurements should be made on isolated cells. d. AFM Cantilever Selection/Coating for Cell Studies AFM cantilevers can vary in their own stiffness and structure. For cell studies, including ECs, cantilevers with a low spring constant (e.g. 0.01-0.1 N/m) with blunt or rounded tips to avoid damaging the plasma membrane are a good choice ( 349 ). One commonly used cantilever for vascular cell AFM is the MLCT from Bruker. Cantilevers can also be coated with proteins such as fibronectin in order to assess cellular adhesion ( 347 ). e. AFM Laser Alignment/Cantilever Calibration After the appropriate cantilever is selected and mounted on the holder of the AFM head, the AFM laser must be aligned to the tip of the cantilever per manufacturer’s protocol. After laser alignment and in order to obtain reliable results, the AFM cantilever sensitivity and spring constant must be calibrated according to the manufacturer’s protocol. This involves indenting the cantilever on an area of very high stiffness (e.g. on the tissue culture plastic or glass that is devoid of cells). f. AFM System Settings for Force Collection Within the AFM software, the following parameters are a good starting point to assess EC stiffness ( 346 ) ( 347 ): force/traveling distance of 1.6 μm, scanning rate of 0.3 Hz, velocity between 800-1,000 nm/s, sampling rate of 625 Hz, and a trigger/set point of 0.3 V. g. Force Curve Acquisition For cells, a good practice is to obtain ñ=30 force curves from the same spot in the cell. For force curve acquisition, consistency is key. Accordingly, measurements should be made at consistent spots on the cells, e.g. halfway between cell boundary and the nucleus. This is important because cell stiffness increases closer to the nucleus and decreases toward the cell boundary ( 350 ). The ~30 force curves should be repeated in n=8-12 cells per dish. If your experimental question involves conditions that are in separate dishes of cells, ideally, acquiring force curves from one dish of cells from each group on the same cantilever will reduce variability. However, this is not always possible as cantilevers can get damaged by the cell or culture debris, which is why calibration is important. h. AFM Analysis Most biological AFM systems have their own software in which to acquire and analyze force curves, typically using the Hertz model or the Oliver-Pharr method (a modified Hertz model) ( 346 )( 347 )( 351 ). For example, the MPF-3D-BIO AFM from Asylum Research uses proprietary software on the backbone of Igor Pro. These software packages can be used to analyze both incremental modulus (Einc), or stiffness, in addition to adhesion if that experimental design is incorporated. In terms of statistics, we contend that the best practice for EC AFM data is as follows: all ~30ish force curves from each of n=8-12 cells per dish should be averaged so that each dish of ECs is a statistical n=1 (assuming that each dish is a separate biological “n”). Then these data can be collated in an appropriate statistical test, depending on the question being asked. This method is the most conservative and will help ensure a true biological difference in mechanical measurements in cells. Doing statistics on the number or force curves or number of cells could artificially conflate the biological meaning of the data. As such, we recommend doing statistics on the number of biological replicates, rather than the number of cells from which curves were obtained. L. Approaches to Assess Endothelial Metabolism Cellular metabolism can be divided into two groups: anabolic metabolism (the pathway that builds molecules by using energy) and catalytic metabolism (the pathway that breaks down large molecules and generally produces energy). Anabolic metabolism primarily occurs in the liver, skeletal muscle, and adipose tissue but is minor in ECs. In this paragraph, we will thus focus on catalytic metabolism in ECs. Major catabolic metabolisms in ECs are glucose metabolism, lipid metabolism, and amino acid metabolism ( Fig. 19 ). Of note, catalytic metabolism without generation of ATP are not discussed in these guidelines. Figure 19. Open in a new tab Schematic diagram of glucose, fatty acid, and glutamine metabolism. FA, fatty Acid; GLUTs, glucose transporters; FATPs, fatty acid transporters; SLCs, solute carrier transporters; CoA, coenzyme A; α-KG, alpha-ketoglutarate. Created in BioRender. Makino, A. (2026) https://BioRender.com/7ue2jjt . Glucose Metabolism in Endothelial Cells: Glucose uptake into the cells is carried out with glucose transporters, including the glucose transporter family (GLUTs) and the sodium-glucose co-transporters (SGLTs). However, due to the lower expression levels of SGLTs, ECs primarily utilize GLUTs ( 352 )( 353 ). Glucose metabolism involves two distinct pathways: glycolysis and oxidative metabolism. Glycolysis occurs in the cytosol, and 2 ATP and 2 NADH are produced during this process. The end-product of glycolysis, pyruvate, will be transported into mitochondria via mitochondrial pyruvate carrier and used for oxidative metabolism. Oxidative metabolism includes citric acid or Krebs cycle and oxidative phosphorylation, and both are carried out in mitochondria and generate 30-32 ATPs per molecule of glucose. Excitable cells (e.g., cardiac myocytes, skeletal muscle cells, and VSMCs) utilize oxidative metabolism due to high ATP demands. However, ECs utilize glycolysis for energy production, accounting for over 85% of total ATP production ( 354 )( 355 )( 356 ). Despite the fact that arterial ECs are constantly exposed to high oxygen levels (~100 mmHg PO 2 ), they generate energy via glycolysis over oxidative metabolism. The reason for endothelial reliance on glycolysis is poorly justified; however, there are several hypotheses: 1) ECs are non-excitable cells; therefore, they simply do not need many ATPs for their survival; 2) ECs avoid utilizing oxidative phosphorylation to minimize excess production of ROS which may lead to oxidative stress that ultimately harms neighboring cells and themselves; 3) ECs use glycolysis because it produces ATPs faster than oxidative metabolism ( 357 ). Low mitochondrial volume density in ECs (~5%) ( 358 ) is also speculated as a reason for endothelial reliance on glycolysis, compared to cardiac myocytes (25-30%). However, it might not be the case since VSMCs, whose mitochondrial density is similar to ECs, generate more energy using oxidative metabolism than glycolysis ( 359 )( 360 ). There are two side branches of glycolysis: the pentose phosphate pathway (PPP) and the hexosamine biosynthesis pathway (HBP). PPP uses glucose 6-phosphate to produce NADPH, a co-factor for the redox system with an antioxidant effect ( 361 )( 362 ). HBP utilizes fructose-6-phosphate (an intermediate in glycolysis), glutamine, acetyl-CoA, and uridine to generate UDP-GlcNAc, a substrate for glycosylation. About 2-3% of glucose is metabolized in the HBP, and UDP-GlcNAc-mediated posttranslational modification plays a fundamental role in protein activation/deactivation ( 363 ). Those two pathways are pivotal in cell physiology and pathophysiology; however, we will not discuss details in these guidelines. The summary of methods used for measuring glucose metabolism is listed in Table 14 . Table 14. Analytical method to assess glucose metabolism Focus Measurement Equipment Advantage Disadvantage ✓ Glucose Uptake 3H-2DG6P Scintillation Counter - High sensitivity - Radioactive handling - No live measurement ✓ Glucose Uptake 2-NBDG Fluorimetry, Microscope, Flow cytometer - Live image - Low sensitivity - May not reflect transport activity correctly ✓ Glucose Uptake Biproduct after 2DG6P conversion by G6PDH Fluorometer Luminometer - Fast and easy - No live measurement - Influenced by endogenous G6PDH activity ✓ Glycolysis End Product Pyruvate, Lactate Plate Reader - Fast and easy - No live measurement ✓ Cell Metabolism Glycolysis Oxidative metabolism FAO- and amino acid-mediated mitochondrial respiration Seahorse XF Extracellular Flux Analyzer - Easy - Measurement in live cells - Open in a new tab Glucose uptake in the cells is the first step of glucose metabolism, and radioactive-labeled substrates (e.g., 3 H-2-deoxyglucose [ 3 H-2DG]) were traditionally used to measure glucose uptake. 2-DG is a glucose analog that can be transported into the cells and phosphorylated but is not metabolized further. Therefore, 3 H-2DG accumulates in the cells, which can be detected by the scintillator ( 364 ). This method gives us the highest sensitivity; however, the handling of radioactive material is a concern. To overcome it, fluorescent glucose tracer 2-[N-(7-Nitrobenz-2-oxa-1,3-diaxol-4-yl) amino]-2-deoxyglucose (2-NBDG) was developed. 2-NBDG can be transported into the cells in the same way as glucose, through GLUTs; however, the transport rate is much slower than for glucose ( 365 ) due partly to the size difference, which is a disadvantage of this method. The advantage of a fluorescent tracer is that glucose uptake can be monitored in a live cell. Other conventional ways to evaluate glycolysis activity are by measuring its byproducts, including pyruvates, lactates, and NADPH ( 366 ). Many assay kits are commercially available for this purpose, and their protocols are relatively easy and fast. However, the limitation is that the samples must be lysed for the assay, so there is no live measurement. Over the past twenty years, the Seahorse XF Extracellular Flux Analyzer (Agilent Technologies, Inc.) has become the “go-to” technology for the measurement glycolysis ( Fig. 20A ). This machine was introduced by Seahorse Bioscience in 2006 and has evolved to change the way cellular metabolism is monitored in live cells. The machine allows for the measurement of multiple metabolic cascades, including extracellular acidification rate (ECAR) ( Fig. 20B ), oxygen consumption rate (OCR) ( Fig. 20C ), proton efflux rate (PER), and ATP production. Using different substrates, we can also determine the lipid- and amino acid-mediated OCR in the cells. Agilent Technologies sells many assay kits designed for different metabolic pathways with relatively simple protocol. There are 24- and 96-well plates for cell plating, and all required reagents are provided in the kit. Minor disadvantages of this system include that 1. cells isolated from tissue using magnetic beads cannot be used because the beads interfere with signal detection, 2. comparing acute hypoxia-exposed cells with controls is challenging due to the required 1-hour incubation under 1% oxygen before the assay begins, and 3. the system is incompatible with buffers containing CO 2 /HCO 3 − , which can disrupt intracellular pH regulation and likely affect metabolic activity. Figure 20. Open in a new tab Seahorse XF Extracellular Flux Analyzer. (A) Schematic diagram of the system. Glycolysis stress test profile. Created in BioRender. Makino, A. (2026) https://BioRender.com/k9haz46 . (B) ECAR, extracellular acidification rate. Cell mito stress test profile. (C) OCR, oxygen consumption rate. Lipid Metabolism in Endothelial Cells: ECs take up fatty acid (FA) from the blood via passive diffusion, using transmembrane proteins, and through transcytosis. FAs absorbed in ECs are utilized in fatty acid oxidation (FAO), transported to the neighboring cells, or used for lipid droplet formation and storage. Short- and medium-chain FAs (< 8 carbon atoms) can enter mitochondria by passive diffusion, whereas the transport of long-chain FA (LCFA) into mitochondria relies on the carnitine shuttle ( 367 ). FA-acyl-CoA synthetases first activate LCFA to LCFA-acyl-CoA, and then carnitine palmitoyl transferase (CPT)1 in the outer mitochondrial membrane converts LCFA-acyl-CoA to acyl-carnitine derivatives. The derivatives cross the inner mitochondrial membrane into the mitochondrial matrix by carnitine-acylcarnitine translocase. They are then transferred back to acyl-CoA by CPT2. Acyl-CoA becomes acetyl-CoA via β-oxidization, utilized in the Krebs cycle for ATP generation. Lipid metabolism is generally not a significant energy source in ECs ( 356 ). Like glucose metabolism, lipid metabolism starts from lipid incorporation in the cells. Therefore, measuring lipid uptake in the cell is one way to assess lipid metabolism. Lipid uptake is used to be evaluated using radiolabeled lipid ( 368 ), but has now been replaced by fluorescent FAs. FAO activity correlates with NADH production. In the presence of iodonitrotetrazolium (INT), NADH generation could be determined by measuring red-colored formazan (a reduced form of INT). This method might be the most effective way to measure lipid metabolism, and there are many commercially available kits for this purpose. Seahorse analyzers can determine FAO-mediated OCR using the Palmitate Oxidation Stress Test Kit. Basically, this kit with glucose/glutamine-depleted media lets cells utilize palmitate as a sole substrate during oxidative metabolism. Using etomoxir (a FAO inhibitor) during the measurement can confirm the FAO-mediated oxygen consumption. Amino Acid Metabolism in Endothelial Cells: Proteins are made from twenty amino acids: four nonessential amino acids, seven conditionally essential amino acids, and nine essential amino acids. Those amino acids are primarily utilized to synthesize protein; however, they are also a substrate for ATP generation ( 369 ). Some amino acids are converted into intermediates of Krebs cycles, generating ATP, while others could become precursors of glycogenesis. Among the 20 amino acids, glutamine is a nonessential amino acid and the most abundant amino acid in the body. Glutamine is transported from the bloodstream into cytosol using the solute carrier (SLC) transporters (e.g., SLC1, SLC6, SLC7, and SLC38), and cytosolic glutamine will then be transferred into the mitochondrial matrix by SLC1A5 and SLC15A transporter family ( 370 ). In the mitochondria, glutaminase converts glutamine to glutamate, and glutamate dehydrogenase converts glutamate into α-ketoglutarate, entering the Krebs cycle. Generally, ATP production in ECs is substantially lower via glutamine metabolism than via glycolysis. The amino acid utility is, however, increased when glucose availability is low (e.g., starvation) ( 371 ). In addition, the reliability of glutamine metabolism varies among cell types and tissues. For example, the corneal ECs produce ~50% of ATP using glutamine as a Krebs cycle intermediate ( 372 ). In human umbilical ECs, glutamine can supply 30% of carbon in the Krebs cycle, comparable to glycolysis and FAO-derived carbon ( 373 ). Radiolabeled and fluorescent amino acids are commonly used for assessing amino acid uptake. There are many assay kits to measure amino acid uptake and cellular glutamine/glutamate concentration. However, there are few methods available to measure the level of amino acid-mediated oxidative metabolism, except with the Seahorse Analyzer. Therefore, there is an urgent need for developing tools for assessing amino acid metabolism. Metabolomics: The technological advancement in omics during the past 20 years was astonishing, and now the cost is getting friendly to investigators. Omics technology enables us to screen and understand the levels and alterations of small molecules within a short time. Metabolomics refers to metabolite profiling, and the data allow us to determine the diverse metabolite levels and their interactions. Traditional metabolomics are carried out with gas chromatography-mass spectrometry (GS-MS), liquid chromatography-MS (LC-MS) ( 374 ), or nuclear magnetic resonance (NMR) spectroscopy ( 375 ). MS is more sensitive than NMR spectroscopy; however, NMR spectroscopy data show higher reproducibility than MS data. Modifying the NMR method, including isotope labeling and hyperpolarization, markedly increases the sensitivity of NMR spectroscopy in detecting the metabolite. There are few differences in methods among glucose, lipid, and amino acid metabolism measurements. In other words, there are not many methods available to evaluate and compare the metabolic cascade. Metabolomics is an excellent tool for screening diverse metabolites; however, it is still limited in accessibility due to the requirement for specific systems and analytical skills. After screening metabolites, the results obtained from omics should be confirmed using targeted commercial metabolite kits, such as NAD + /NADH, ATP, lactate, glutathione (GSH/GSSG), TCA-cycle intermediates, or amino-acid quantification kits, or by functional metabolic assays using a Seahorse analyzer. These targeted approaches provide pathway-specific validation of the changes identified in the initial metabolomics screen; however, there are limitations in some of the metabolites to be re-evaluated with conventional methods. Therefore, improvements in metabolic assays are required to continue to move this field forward. In addition, the live imaging of endothelial metabolism needs improvements, since none of the methods can give sufficient resolution to identify cellular metabolism specific to ECs in vivo . 3. PHARMACOLOGICAL MEASUREMENTS Pharmacological Agents Used to Assess Endothelial Function For the last decades, many pharmacological agents have been used to inhibit major sources of NO, prostanoids, EDH and ROS, to assess their contribution in endothelial (dys)function in varied situations. In this section, we provide an overview of the most widely used inhibitors of these mediators. For many of these agents, selectivity has been proven, however, for others, off-targets effects, often associated with different doses or chronic vs acute administration, cannot be ruled out. Then, whenever possible, confirming inhibition (or activation), testing several compounds, or using genetically modified models, would be a wise approach to assure selectivity of the observed effects of the pharmacological agents used. Although the pharmacological agents discussed below are commonly used in ex vivo myography, the pharmacological principles discussed in this section are broadly applicable across multiple experimental platforms. It is important to note, however, that specialized techniques often require distinct experimental conditions that influence how, when, or whether pharmacological agents can be applied. For instance, patch-clamp electrophysiology relies on highly controlled internal and external solutions (e.g., precise pH, absence of albumin, defined ionic composition), and the sensitivity of the equipment may preclude the addition of drugs during active recordings. In these situations, treatments may need to be performed prior to the recording period or adapted to the specific constraints of the protocol. Similarly, AFM measurements and certain en face preparations may involve mechanically constrained environments and/or fixed tissues, limiting the feasibility of introducing pharmacological agents during data acquisition. Thus, while the underlying signaling pathways remain consistent, the timing, method of application, and allowable concentrations of pharmacological agents can differ markedly across techniques. These differences may alter time-dependent responses or concentration-effect relationships and therefore require careful interpretation when comparing pharmacological outcomes across diverse methodological approaches. Pharmacological Agents Used for Investigating the Role of NO Nitric oxide can be released from different cell types from the three isoforms of NO synthase (NOS): neuronal NOS (nNOS or NOS1), inducible NOS (iNOS or NOS2) and endothelial NOS (eNOS or NOS3), that use L-arginine and molecular oxygen as substrates as well as several cofactors such as reduced nicotinamide-adenine-dinucleotide phosphate (NADPH), flavin adenine dinucleotide (FAD), flavin mononucleotide (FMN), and (6R-)5,6,7,8-tetrahydrobiopterin (BH4). In addition, all NOS bind calmodulin and contain haem/heme. nNOS and eNOS are constitutively expressed in central and peripheral neurons and some other cell types and in ECs respectively, and generally also depend on Ca 2+ . iNOS is induced in many cell types in response to inflammatory molecules such as lipopolysaccharide and cytokines, but also by several other factors, and generates large amounts of NO in a Ca 2+ -independent manner ( 376 ). nNOS and mainly eNOS are involved in the control of vascular tone and vasodilation and blood pressure regulation, among many other functions. However, different cardiovascular risk factors can promote eNOS uncoupling due, for example, to L-arginine or BH4 deficiency where superoxide anion (O 2 .- ) rather than NO is produced, being one of the major causes of endothelial dysfunction ( 377 ). Moreover, low-grade inflammation is present in most cardiovascular diseases with augmented levels of local and circulating proinflammatory molecules that can upregulate iNOS that, in a context of excessive ROS generation (aka oxidative stress), can also contribute to endothelial dysfunction through the production of the oxidative nitrogen species peroxynitrite (ONOO − ) ( 376 ). Because all NOS use L-arginine as substrate, pharmacological blockade of NOS(s) has been mostly performed using L-arginine analogues ( Table 15 ) to identify the role of NO in vascular responses. However, these compounds lack selectivity for specific isoforms and additional compounds were synthesized mainly with the aim of inhibiting pathological iNOS without affecting constitutive eNOS and nNOS and identifying the specific contribution of this pathological NOS in endothelial dysfunction in different conditions. Table 15. Common pharmacological agents used to assess the role of NO, COX and ROS in endothelial function Pathway Target Pharmacological agent Mechanism of action Off-target effects Reference NO All NOS L-NAME L-arginine analogues that inhibit NO production by NOS from L-arginine NO release from the nitro group via Fenton reaction in long term applications ( 391 ) L-NOARG - L-NMMA - iNOS 1400W Highly selective iNOS inhibitor competing with L-arginine Very little degree of other NOSs inhibition ( 392 ) S-methylisothiourea Competitive inhibitor of iNOS activity at the L-arginine site Some degree of eNOS activity inhibition ( 393 ) Prostanoids All COXs Indomethacin (and other NSAIDs) Inhibition of COX activity Nonselective inhibition of COX-1 and COX-2 COX-1 SC-560 Highly selective COX-1 inhibitor It might act in some cell types as an unselective COX inhibitor ( 394 ) ( 395 ) COX-2 NS398 Highly selective COX-2 inhibitors Non selectivity at high doses ( 396 ) ( 397 ) COXIBs DUP697 DFU PGIS Tranylcypromine Prostaglandin I2 synthase inhibitor Monoamine oxidase inhibition TXAS Ozagrel Selective thromboxane A2 synthase inhibitor PGES Compound III Selective microsomal prostaglandin E synthase (mPGES)-1 inhibitors Partial inhibitory activity of L-PGDS at high concentration ( 398 ) Compound 118 Weak to moderate inhibition of mPGES-2 ( 399 ) IP RO1138452 RO3244794 CAY10441 Structurally distinct IP receptor antagonists Not expected if used at recommended concentrations ( 400 ) ( 381 ) TP SQ-29548 S18886 L-655,240 GR 32191 Structurally distinct selective TP receptor antagonists ( 381 ) EP1 SC19220 SC51322 ONO-8130 Selective EP1 antagonists ( 381 ) ( 401 ) EP2 PF-04418948 Selective EP2 antagonist EP3 L-798106 Selective EP3 antagonists EP4 GW627368X Selective EP4 antagonists Reactive Oxygen Species O 2 .− (PEG)-SOD (Cell permeable)-superoxide dismutase that catalyses the conversion of O 2 .− to H 2 O 2 . Tempol Redox cycling agent (catalyses the dismutation of O 2 .− , facilitates catalase-like metabolism of H 2 O 2 and prevents Fenton signalling and ·OH formation) - Modulation of K + channels - Inhibition of the sympathetic nervous system ( 402 ) ( 403 ) NAC (i) reductant of disulfide bonds, (ii) scavenger of reactive oxygen species and/or (iii) precursor for glutathione biosynthesis - Elevations of H 2 S and sulfane sulfur species inside cells. - Antiinflammatory effects. - Inhibits ER stress ( 404 ) ( 405 ) H 2 O 2 (PEG)-Catalase (Cell permeable)-catalase that converts H 2 O 2 into H 2 O Several Nox(s) DPI Flavoprotein inhibitor Inhibition of other enzymes besides Nox(s) ( 406 ) ( 407 ) ( 389 ) Apocynin Initially thought to interfere with the intracellular translocation of the cytosolic p47phox and p67phox subunits to the membrane - Direct scavenger of ROS rather than Nox inhibitor - Additional targets (RhoK, PI3K/Akt) VAS2870 Covalent binding to Cys at Nox(s) active sites - Alkylate thiols - Modulates PKC signalling Nox1 ML-171 Nox1 inhibitor Inhibits other Nox(s) ( 389 ) ( 385 ) GKT137831 Nox1/Nox4 inhibitor and partial Nox5 inhibitor - Inhibits several Nox(s) - Oxidant scavenging effects NoxA1ds Inhibits interaction of NoxA1 with Nox1 of the active, canonical Nox1 isozyme ( 408 ) Nox2 Nox2-dstat Inhibits binding of the organizer cytosolic subunit p47phox to loop B of Nox2 ( 409 ) Nox4 GKT137831 Nox1/Nox4 inhibitor and partial Nox5 inhibitor - Inhibits several Nox(s) - Oxidant scavenging effects ( 385 ) ( 389 ) Nox5 ML090 Nox5 inhibitor Inhibits other Nox(s) (388) XO Allopurinol Inhibits xanthine oxidase activity and ROS produced during the purine degradation pathway Activates Nrf2 ( 410 ) Mitochondria Mito-Tempo Mitochondrial-targeted SOD mimetic ( 411 ) Mito-Q Mitochondria-targeted antioxidant derived from ubiquinone (Coenzyme Q10) Inhibits NF-kB ( 412 ) ER stress PBA TUDCA Chemical chaperones that stabilize protein conformation, improve ER folding capacity, and inhibit ROS formation during ER stress Multiple molecular targets within the ER ( 405 ) ( 413 ) GPx Ebselen Mimic of glutathione peroxidase - Inhibits some Nox(s) - Effects on many additional targets ( 389 ) Nrf2 Dimethyl fumarate Electrophilic compounds that bind cysteine 151 in KEAP1 and prevent Nrf2 degradation - KEAP1-mediated NRF2-independent actions - Action on other KEAP1 partners different from Nrf2 ( 390 ) ( 414 ) L-sulforaphane Bardoloxone Open in a new tab Note: This list is not intended to be exhaustive or to include all existing drugs targeting the specific pathways. Rather, it provides an overview of selected compounds that have demonstrated endothelial effects under various experimental conditions. The listed references primarily address issues of compound selectivity, not their endothelial effects. No references are provided for drugs already in clinical use or for compounds that mimic natural enzymes. COX: cyclooxygenase; DPI: diphenyleneiodonium; ER: endoplasmic reticulum; GPx: glutathione peroxidase; L-NOARG: NG-nitro-L-arginine; L-NMMA: NG-monomethyl-L-arginine; L-NAME: NG-nitro-L-arginine methyl ester; NAC: N-acetyl cysteine; NOS: NO synthase; Nox: NADPH oxidase; Nrf2: Nuclear factor-erythroid 2-related factor-2; NSAIDs: non-steroidal anti-inflammatory drugs; PBA: 4-Phenylbutyric acid; PEG: Polyethylene glycol; PG: prostaglandin; ROS: reactive oxygen species; SOD: superoxide dismutase; TUDCA: tauroursodeoxycholic acid; XO: xanthine oxidase. Pharmacological Agents Used for Investigating the Role of COX-Derived Products Prostanoids are lipid mediators enzymatically synthesized from the polyunsaturated omega-6 fatty acid arachidonic acid that is released from membrane phospholipids by the action of phospholipase A 2 upon physiological or pathological stimulation. COX-1 (a constitutive isoform) and COX-2 (induced in response to varied stimuli) convert arachidonic acid into the endoperoxides prostaglandin (PG) G 2 and PGH 2 . This PGH 2 is further isomerized to each prostanoid by the corresponding synthases: PGE synthase (mainly microsomal mPGES-1) for PGE 2 , PGI synthase (PGIS) for PGI 2 (or prostacyclin), thromboxane synthase (TXS) for TxA2, PGD synthases (PGDS) for PGD 2 , and PGF synthase (PGFS) for PGF 2 a. Prostanoids mediate their effects trough specific G-protein-coupled receptors that are differentially expressed in many tissues and cells, triggering varied downstream signaling in physiological and pathological conditions. Specifically, PGI 2 activates the IP receptor, TxA 2 binds the TP receptor, PGE 2 has 4 receptor subtypes (EP1-4), PGD 2 binds to 2 receptors (DP1-2) and PGF 2 a binds the FP receptor ( 378 ). COXs, isomerases and prostanoid receptors are expressed at the vascular level in different cell types ( 378 ). In fact, it is well known that apart from NO, ECs release PGI 2 in physiological conditions. In addition, ECs can also release endothelium-derived-contracting factors (EDCF) ( 379 ) which in many circumstances belong to the family of COX-derived factors. Experimentally, endothelium-dependent COX-dependent contractions are appreciated in response to ACh and other vasoactive substances such as arachidonic acid or the Ca 2+ ionophore A23187 ( 379 ). These EDCFs are particularly evident in disease conditions such as obesity, hypertension, aging and many others, but they can also occur in arteries from healthy subjects ( 379 ). The specific EDCF highly depends on the animal species, vascular bed and particular disease ( 379 ). A role for constitutive COX-1 has been described for EDCF production. However, the inducible COX-2 isoform, which is upregulated in several cardiovascular diseases, likely as a reflection of the proinflammatory phenotype, has also been described as a source of EDCF in both animal models and humans. Among specific prostanoids, TxA2, PGF 2 α, PGE 2 , and even PGI 2 have been shown to be involved in endothelial dysfunction by modulating the production or effects of vasodilator factors, contractility and oxidative stress and inflammation ( 379 )( 380 ). Over the last decades, numerous studies have tried to uncover the specific COX and/or isomerase or prostanoid involved in endothelial (dys)function using dozens of pharmacological inhibitors for these enzymes or receptors ( Table 15 ) ( 381 )( 382 )( 383 ). Some drugs show greater selectivity for COX-1 or COX-2. Specific inhibitors for some isomerases and receptors are also available. However, it is also important to recognize that in some conditions, specific receptors can bind several prostanoids (i.e., TP receptors can bind TxA 2 , PGF 2 a and PGI 2 ) ( 384 ) which might complicate interpretation of the results. In this context, performing complementary experiments and controls is recommended for a more accurate interpretation of results based on these pharmacological agents. Pharmacological Agents Used for Investigating the Role of ROS ROS are reactive derivatives of oxygen metabolism. These include highly unstable short half-life molecules with unpaired electrons such as O 2 .− and hydroxyl radical ( . OH). The majority of O 2 .− generated is rapidly converted to the non-radical ROS H 2 O 2 which is more stable with longer half-life. This can be achieved spontaneously or by superoxide dismutase (SOD) enzymes, such as cytosolic Cu/Zn-SOD (SOD1), mitochondrial Mn-SOD (SOD2) and extracellular EC-SOD (SOD3). Additionally, O 2 .− can react with NO forming and ONOO − . H 2 O 2 may be converted into water by the enzyme catalase or other enzymatic systems such as glutathione peroxidase-1 or the thioredoxin (TRX) system ( 385 ). O 2 .− is formed by the univalent reduction of molecular oxygen mediated by enzymes such as NADPH oxidases (Nox), xanthine oxidase (XO), lipoxygenase, COX, CYP450 isoforms, monoxygenases and uncoupled eNOS, and can also be generated non-enzymatically by the mitochondrial electron transport chain, the endoplasmic reticulum (ER), and peroxisomes ( 385 ). The major source of ROS at the vascular level is the Nox family, which is upregulated in many cardiovascular diseases ( 385 ). There are seven Nox isoforms of which Nox1, Nox2, Nox4 and Nox5, are expressed in fibroblasts, ECs and VSMCs, and are regulated by a wide variety of stimuli including elevated intravascular pressure, proinflammatory cytokines and vasoactive agents such as endothelin-1 and angiotensin II, among others. Of those, Nox5 is only expressed in humans. Nox(s) are professional O 2 .− generating enzymes. Moreover, some types of Nox including Nox-4, can directly produce H 2 O 2 . Many preclinical studies performed in different vascular beds from animal models and studies using human arteries have shown that various antioxidants and inhibitors of ROS production improve endothelial function in different cardiovascular diseases. In this sense, there is a vast variety of compounds that have been used to inhibit ROS sources, activate their detoxification or directly scavenge ROS. A complete description of the very many compounds available is not the scope of these guidelines and there are focused reviews on the topic ( 386 )( 387 ). Here we show some of the most widely used inhibitors for evaluation of the role of ROS in endothelial function ( Table 15 ). In general, most synthetic compounds inhibit ROS availability but most of them lack specificity for one single ROS or source. For example, many Nox inhibitors have been designed or are currently under development ( 386 )( 388 ). However, the specificity of Nox inhibitors remains controversial, and researchers should be careful on aspects such as concentrations used in vitro or in vivo before assuming that effects of single inhibitors unambiguously identify contribution of specific Nox ( 389 )( 390 ). Moreover, additional off-target effects have been described for several compounds that could contribute to vascular tone modulation, and this should be taken into account when interpreting experimental data. Use of several compounds and controls and appropriate quantification of specific ROS would enhance the quality of the obtained finding ( 389 ). Increased levels of ROS activate nuclear factor (erythroid-derived 2)-like 2 (Nrf2), a master regulator of the antioxidant response, which is activated to counteract oxidative stress. Nrf2 controls the expression of about 250 genes including those encoding antioxidant enzymes such as those involved in glutathione and TRX systems, SOD, catalase, and hemoxygenase-1, among many others ( 391 ). Interestingly, activation of Nrf2 prevents endothelial dysfunction in several animal models. Among these activators are electrophilic compounds that bind KEAP1 (an Nrf2 adaptor that targets Nrf2 for ubiquitination and proteasomal degradation) that prevents its ability to target Nrf2 for degradation, leading to Nrf2 accumulation and upregulation of a large network of cytoprotective proteins ( Table 15 ). Endothelium-dependent hyperpolarization (EDH) Endothelial cell hyperpolarization (EDH) is generated by the activation of two types of Ca 2+ -activated potassium channels found in the endothelium of arteries, K Ca 3.1 (aka IK Ca ) and K Ca 2.3 (aka SK Ca ). EDH is a particularly important vasodilator mechanism in small arteries, as compared with larger arteries their VSMCs have a greater density of voltage gated Ca 2+ channels (VGCC). As vasodilation follows a reduction in VGCC open probability, small arteries are very sensitive to the influence of hyperpolarization. EDH spreads to the adjacent VSMCs by two parallel mechanisms, passive spread via myoendothelial gap junctions and the release of diffusible factors. The latter includes the K + ions that efflux through endothelial IK Ca and SK Ca channels, which causes vasodilation by recruiting K IR channels and activating Na + /K + -ATPase. Under defined, often pathological, conditions other factors such as EETs and H 2 O 2 can be generated and act as an EDH by activating K Ca 1.1 (BK Ca ) channels present in the VSMCs ( 416 ). Experimental design must also consider the possibility that NO has significant input as a hyperpolarizing factor, mainly via VSMC K Ca 1.1. Because endothelial K Ca channels underpin EDH, any maneuver that raises cytoplasmic Ca 2+ concentration will cause EDH-mediated vasodilation. Normally, EDH works alongside NO and, in a small number of arteries, PGI 2 . It can sustain vasorelaxation if these vasodilator pathways are compromised, for example by disease ( 417 ). How to study EDH: EDH can be assessed in isolated arteries either directly, by measuring VSMC and/or EC membrane potential with microelectrodes or indirectly by measuring vasorelaxation in a wire or pressure myograph ( 1 ). In both cases, other endothelial vasodilator pathways must be blocked to isolate and quantify EDH effects. This can be achieved using NO synthase inhibitors such as L-NAME or L-nitroarginine and a COX inhibitor such as indomethacin. In most healthy arteries, blocking COX has little or no effect and can be discontinued if this is shown to be the case in control experiments. Below, you will find our recommended experimental protocol, conditions, and pharmacological approaches to evaluate EDH. a. Direct Microelectrode Measurement of EDH: This is the most desirable approach to study EDH, but experimentally more demanding than simply measuring tension change reflecting the action of EDH ( 418 ). As described above, the most widely used agonist for activating endothelium is ACh. In most arteries, ACh-induced vasodilation is mediated primarily by endothelial M 3 receptors, which raise cytosolic Ca 2+ and activate EDH pathways. Evidence for functional M 3 receptors on VSMC is limited. The first step in an experimental protocol should be to assess the overall integrity of the endothelium, by measuring its ability to cause vasorelaxation in arteries pre-constricted with an agonist, such as phenylephrine, or generating spontaneous, myogenic constriction. A vasorelaxation of >75% in mouse arteries and >90% in rat arteries is considered indicative of a functional endothelium in resistance arteries. Importantly, these values can vary greatly depending on multiple factors, including how the concentration of the pre-constricting agent is determined (e.g., percentage of maximal response), the type of vessel studied, and whether the preparation is from an animal model or human tissue; therefore, this value should be interpreted with these considerations in mind. The next step is to block NO synthase and, if required, COX, and impale an individual VSMC to record membrane potential. EDH can then be stimulated by cumulative or non-cumulative addition of, for example ACh. Resting potentials vary somewhat between arteries but are usually around −45 to −55mV. EDH will hyperpolarize the cells to membrane potentials close to E K , i.e., circa −70 mV. Technically more demanding, but more informative, are membrane potential recordings obtained during either agonist induced or myogenic vasoconstriction. Here simultaneous recording of hyperpolarization (both repolarization and hyperpolarization beyond the original resting potential) and vasorelaxation due to EDH enables the two to be linked. When designing experiments to probe EDH, consideration must also be given to the fact that the EDH profile can be altered during vasoconstriction. In rat mesenteric small arteries, EDH induced by increasing concentrations of ACh is abolished by block of SK Ca channels using apamin alone, while in the same arteries undergoing depolarization and vasoconstriction to phenylephrine, block of both SK Ca and IK Ca channels is necessary to abolish both the hyperpolarization and tension change due to EDH ( 419 ). Finally, although tricky, it is possible to record EC hyperpolarization directly, and this can be useful in some experimental settings. Although this has been reported using arterial segments that have been cut open and pinned out, this approach is not recommended as it completely changes the tension experienced by the cells. It is possible to align a small side branch uppermost when mounting an artery segment in a wire myograph. Cutting off the side branch then leaves a small ‘window’ to provide access for a microelectrode, the first contact and impalement being in an EC ( 420 ). Alternatively, EC membrane potentials can be recorded from inverted arteries ( 421 ). In any case, confirming impalement of EC rather than other mural cells is recommended, for instance, by including a dye in the electrode solution ( 147 )( 421 )( 422 ). b. Indirect Assessment of EDH by Vasorelaxation: Most published EDH data have been obtained in this way. The approach is similar to “a” but without microelectrodes. As vasorelaxation is a proxy for EDH, arteries must be partially constricted. Inhibition of vasorelaxation when vasoconstriction is induced by Krebs buffer containing elevated (isosmotic substitution, usually >45 mM) levels of K + is necessary to show that hyperpolarization is responsible. c. The Extent of Background Vasoconstriction is Important: When designing experiments, it is also important to consider any possibility of functional antagonism, which may occur with maximal or near maximal background vasoconstriction. In mesenteric arteries, as depolarization and contraction to phenylephrine increases, functional antagonism depresses the ability of exogenous K + to evoke hyperpolarization and vasorelaxation. As EDH responses were not depressed, this observation was used to argue against K + as a diffusible EDH. However, with maximal vasoconstriction the effect of K + is right-shifted to higher concentrations. This functional antagonism reflects the intercellular accumulation of K + exiting VSMCs during vasoconstriction through BK Ca channels and saturating the hyperpolarizing ability of exogenous K + . The effect is reversed during submaximal vasoconstriction or by the addition of iberiotoxin to block VSMC BK Ca ( 423 )( 424 ). d. Choice of Agonist to Stimulate Background Constriction is Important: Appropriate control experiments are also necessary to ensure the profile of EDH (both hyperpolarization and vasorelaxation) is not influenced by the particular agonist used to stimulate background vasoconstriction. In human coronary arteries, endothelium-dependent reversal of vasoconstriction to endothelin is mediated by the hyperpolarizing action of H 2 O 2 ; but if the thromboxane-mimetic U46619 is used as a constrictor, endothelial release of NO explains vasorelaxation ( 425 )( 426 ). Furthermore, repeated stimulation with U46619, but not phenylephrine, will selectively abolish SK Ca channel input to EDH ( 427 ) e. Agents Used to Block EDH and Associated Vasodilation: Classically, the EDH pathway is defined by a sensitivity to simultaneous block of endothelial K Ca 2.3 and K Ca 3.1 channels, but resistance to block of K Ca 1.1 channels (using the specific toxin, iberiotoxin). This approach is important, as it effectively removes EDH at source. As EDH spreads in part via myoendothelial gap junctions, which are very difficult to block without affecting homocellular gap-junctions, this is very useful. In nanomolar concentrations, apamin will provide an effective, selective block of K Ca 2.3 and historically was used in combination with charybdotoxin to block K Ca 3.1 channels. However, charybdotoxin is non-selective and its use has been superseded by selective agents such as TRAM-34, a derivative of the anti-fungal agent clotrimazole, and the benzothiazinone derivative NS6180. Alternatively, UCL 1684 can be used as a potent blocker of all small-conductance K Ca channels (K Ca 2.1-2.3), offering a broader inhibition when the contribution of all SK subtypes is of interest. In low micromolar concentrations each selectively blocks K Ca 3.1 in isolated arteries. Both positive and negative gating modulators are also available for studying K Ca 2.3 and K Ca 3.1 channel input to EDH, but even with benzathiazoles such as SKA-31, purported to be a selective activator of K Ca 3.1 channels, caution is needed, and experimental use should include appropriate controls to detect/inhibit activation of K Ca 2.3 and/or K Ca 1.1 ( 428 ). Commonly Used Pharmacological Agents for Evaluating Endothelial Function and Their Mechanisms of Action One of the most essential tools for evaluating vascular function, particularly endothelial function, in laboratory settings is the generation of concentration–response curves. This technique involves exposing isolated blood vessels, maintained in organ baths or myographs, to various agonists (please see section above “ The Assessment of endothelial function ” and reference ( 1 ). These agonists specifically bind to receptors located on ECs and/or VSMCs, triggering responses that result in either vasodilation or vasoconstriction. To assess endothelial function, vasorelaxation and -contraction responses to selected agonists are examined. A diminished relaxation response and/or an enhanced contraction response may indicate endothelial dysfunction and overall impairment of vascular health. To further explore the mechanisms underlying these responses, specific antagonists are employed to inhibit particular pathways, allowing for detailed characterization of signaling routes involved in endothelial and vascular function. The contributing mechanisms and degree of endothelial dependent vasodilation differ across species and vascular beds. While some agents activate similar signaling cascades in preclinical models or the human circulation ( 429 ), others show distinct species and vascular bed specificity. For example, bradykinin-induced vasodilation varies by species and vascular bed, ranging from NO-dependence in rat cerebral arteries ( 430 ) to H 2 O 2 -mediated effects in swine coronary arteries ( 431 ). In humans, swine, and canines, adenosine typically mediates vasodilation through NO pathways while in bovine vessels, the mechanism shifts to involve cyclic AMP (cAMP) instead of NO. Additionally, in some species, adenosine can even cause endothelial-independent dilation ( 432 )( 433 ). The effect of endothelin (ET-1), one of the most potent vasoconstrictors identified to date, can also be studied on arteries in a wire and pressure myographs. ET1 has two receptor subtypes expressed on VSMC, ETA and ETB. In the context of regulating vascular tone, activation of ETA receptors causes vasoconstriction, and activation of ETB receptors may cause vascular relaxation or constriction. To elucidate the effect of ET-1 on an arterial preparation, exogenous ET-1 from 1 pM to 100 nM can be applied at baseline to generate a concentration-dependent contraction. Prior application of ET receptor antagonists 30 mins before the ET-1 concentration contraction curve can determine the relative contribution of the receptor subtypes to the observed constriction. It is important to note that ET receptor antagonists, when applied after ET-1 to the bath, will only cause a partial reversal of the contractile response, and upon washout the ET-1-mediated vasoconstriction rapidly recovers. Thus, ET-1/ETA complexes are not dissociated by the ET-1 antagonists, thereby making it essential to study the effect of the antagonists by pre-applying them to the bath ( 434 ). BQ-123 is a selective antagonist for the ETA receptor antagonist (range ~10-100 nM), but will block ETB receptors in the low micromolar range. BQ-788 will selectively antagonize the ETB receptor and should be used at ~100 nM-1 μM. The non-selective ET receptor antagonist, bosentan (1 μM) will determine the overall effect of ET-1 on vascular tone. When researching ET-1, it is important to consider the tight binding of ET-1 on ETA, such that functional antagonism by many different vasodilators will only transiently reduce but not terminate vasoconstrictions mediated by ET-1-ETA activation ( 434 ). In order to remove ET-1 and terminate the ET-1-ETA activation, CGRP (100 nM) can be applied to promote dissociation of ET-1/ETA-receptor complexes ( 434 ). Application of CGRP will cause the arterial segment to relax and after 10 mins, washout will enable the segment to return to its resting baseline. These examples highlight the complexity and variability of physiological mechanisms of endothelial dependent dilation and are discussed in depth elsewhere ( 435 )( 436 ). Table 16 summarizes key mechanisms and pharmacological means to stimulate endothelial mediated dilation and highlights underlying mechanisms/pathways involved. Table 16 also provides the agonists or agents used to induce vasoconstriction (preconstrict the arteries prior to inducing vasodilation), the receptor types targeted, the associated intracellular signaling mechanisms, and the expected vascular responses linked to each agonist, with a particular focus on endothelial-mediated effects, except for sodium nitroprusside (which is primarily used to assess endothelium-independent relaxation or to distinguish endothelial from VSMC contributions to vasodilation). Table 16: Key agents used in concentration-response curves for studying vascular function, and underlying mechanisms/pathways involved. Agent Receptor Vascular Signaling pathway Expected Response ( arteries from healthy animals/human ) References VASODILATORS Common Pharmacological Agents Used to Induce Vasodilation via the Endothelium or Released by the Endothelium Acetylcholine (ACh) M3 > M2-muscarinic receptors • Gq/PLC/ IP 3 , increase intracellular Ca 2+ , activates eNOS, NO release • EDH activation • Prostacyclin release • Vasodilation ( 2 )( 71 ) ( 436 )( 437 ) Other muscarinic receptor agonists (Methacholine and Pilocarpine) M3 receptor ( 438 ) ( 439 ) ( 439 ) Isoproterenol β 2 adrenergic receptor Gs/PKA/cAMP • Vasodilation ( 440 ) Prostacyclin (PGI 2 ) Prostacyclin receptor (IP) Adenylyl cyclase/cAMP • Vasodilation ( 441 ) Prostaglandin E2 (PGE 2 ) EP4>EP2 Adenylyl cyclase/cAMP • Prostaglandins of the E series (PGEs): These generally dilate arterioles, metaarterioles, precapillaries, and venules in most organs ( 441 ) CYP450 Metabolites (EETs, HETEs) • Activation of -conductance Ca 2+ -activated potassium (BK Ca ) • EETs can increase NO production and/or prostacyclin (PGI 2 ) synthesis in some contexts • EETs and HETEs - Vasodilators in pulmonary circulation • EETs vasodilator; HETEs vasoconstrictor in coronary and peripheral circulation ( 442 ) H 2 S • ROS scavenging • Regulates redox balance • Phosphatidylinositol 3-kinase (PI3K)/Akt pathway activation • anti-inflammatory pathway activation (NFkB, Nrf2) • Activates K ATP channels • Increases NO bioavailability • Stabilizes electron transport chain • Dilation in coronary and peripheral vessels • Dual effects (constriction and dilation) in pulmonary circulation ( 443 )( 444 ) Bradykinin B2 (constitutive) and B1 (inducible) kinin receptor • G αq /PLC/ERK1-2/eNOS signaling, NO release • Releases Prostaglandin • Dilation in coronary and peripheral vessels •Constriction and dilation in pulmonary circulation •Increased glomerular filtration rate in kidneys ( 445 )( 446 )( 447 ) Histamine H1- H4 • Activation of NO and Prostacyclin signaling • Increases vascular permeability • Dilation in coronary and peripheral vessels (H1 and H2 receptors) • Dual effects (Constriction and dilation) in pulmonary circulation ( 448 ) Serotonin 5-HT1 and/or 5-HT2 • 5HT (different sub forms) mediated release of NO; increase in cAMP or Prostacyclin • Depending on cell type expression counteracts dilation (vasoconstriction) Serotonin predominantly induces vasoconstriction across most vascular beds, with some exceptions where it can induce vasodilation, such as in human umbilical arteries and certain microvascular beds ( 449 )( 450 ) Ca 2+ Ionophores (e.g., A23187 aka Calcimycin or Ionomycin) • Increased Ca 2+ flux • Ca 2+ /calmodulin-dependent protein kinases (CaMKs) and phosphatases • Phospholipase C • Enhance Ca 2+ influx in coronary and peripheral circulation • Can lead to constriction and dilation in pulmonary circulation • Can cause endothelial independent dilation in pulmonary and renal circulation ( 451 ) Substance P Neurokinin-1 receptor (NK1R) • Neurokinin 1 (NK1) receptor signaling • Increased NO and prostacyclin production • Triggers histamine release • Increases vascular permeability • Vasodilator in coronary circulation • Dual effects (constriction and dilation) in pulmonary circulation •Different expression of receptor (NK1) between rodents and humans ( 452 ) Amino Acids (e.g. Arginine, Lysine, Tryptophan) • NO mediated activation of cAMP/cGMP • Increase Serotonin • Ca 2+ modulation • cis-WOOH (TRY metabolite via IDO 1 • Generally, cause vasodilation but downstream signaling molecules (e.g. serotonin) can have dual effects ( 453 ) Adenosine Di or Triphosphate (ATP/ADP) Purinergic receptor activation • Triggers NO release • EDH activation • Dilation in coronary and peripheral vessels •Constriction in pulmonary circulation ( 454 ) Sodium Nitroprusside • NO donor Vasodilation ( 455 ) VASOCONSTRICTORS Common Pharmacological Agents Used to Induce Arterial Constriction to Evaluate Endothelial Function U46619 Thromboxane A 2 receptor • Activates phospholipase A 2, via Gq coupling and Rho-kinase pathway, increasing intracellular Ca 2+ • Vasoconstriction ( 456 )( 457 ) Noradrenaline/ Norepinephrine α 1 - and α 2 -adrenoreceptor •G q receptor/PLC/IP 3 /DAG/PKC; PLA 2 •RhoA/ROCK Pathway and Ca 2+ -Independent Contraction • Vasoconstriction ( 458 ) Phenylephrine α 1 -adrenoreceptor • Activates PKC, via G q receptor • Vasoconstriction ( 459 ) Serotonin 5-HT 1 and/or 5-HT 2 • 5-HT 1 : G i/o -inhibition cAMP formation • 5-HT 2 : Gq/11 activation of IP3/PKC/cytosolic Ca 2+ signaling pathway in SMC • Vasoconstriction ( 460 ) ( 461 ) Angiotensin II AT1> AT2 receptors • Gq/PLC,PLD,PLA2/DAG, MAP kinases, tyrosine kinases, tyrosine phosphatases, and RhoA/Rho kinase • Vasoconstriction ( 462 )( 463 ) Endothelin-1 ET A > ET B receptors • Gq receptor/PLC/IP 3 /DAG/PKC/intracellular Ca 2+ • Vasoconstriction ( 464 )( 465 ) PGF 2 α (Prostaglandin F 2 α) FP receptor • Increased intracellular Ca 2+ • Vasoconstriction ( 466 ) Open in a new tab Interpretation of Pharmacological Responses for Ex Vivo Vascular Function Potency (EC 50 ) and/or maximum response (E max ) calculated from concentration-response curves to ACh and other pharmacological agents (see Table 16 ) are key pharmacological parameters for evaluating and comparing endothelium-dependent relaxation responses ( 439 ). For example, in pharmacology, the EC 50 is one of the most widely used parameters to quantify an agonist’s potency and the sensitivity of a tissue to that agonist. For endothelium-dependent vasodilators such as ACh or bradykinin, changes in EC 50 directly reflect endothelial function because relaxation requires endothelial receptor activation, intracellular Ca 2+ mobilization, and subsequent NO/EDH production. A rightward shift (higher EC 50 ) therefore indicates reduced endothelial sensitivity to the agonist, whereas a leftward shift (lower EC 50 ) reflects enhanced endothelial responsiveness. In addition, the use of specific agonists, inhibitors, and pathway activators helps define which endothelial mechanisms, such as receptor signaling, Ca 2+ handling, eNOS activity, or EDH pathways, are responsible for the observed changes in EC50. (for further information please see ( 1 ). The area under the curve (AUC) can also serve as a useful tool for evaluating the participation of endothelial pathways/factors ( 468 ). The difference in AUC (dAUC) between treated (e.g., exposed to a specific endothelium-derived factor inhibitor, please see Table 15 ) and control concentration-response curves ( 468 ) can be calculated for comparative analysis. In some vessels and/or pathological conditions, ACh can evoke contraction at higher concentrations. This was first demonstrated in aorta of spontaneously hypertensive rats (SHR) precontracted with noradrenaline, where ACh at concentrations below 10 −6 M induced relaxation but switched to contraction at higher concentrations ( 469 )( 470 )( 471 ). A similar biphasic response has been observed in mesenteric resistance arteries of hypertensive rats ( 472 )( 473 ) ( Fig. 21 ). Figure 21. Open in a new tab Concentration-response curves to acetylcholine in mesenteric resistance arteries from (A) DOCA-salt hypertensive rats (DOCA), (B) spontaneously hypertensive rats (SHR), and normotensive male Wistar rats (NT). Acetylcholine induced a biphasic response in arteries of *p<0.05 vs. NT (2-way-ANOVA). Image taken with permission from ( 473 ). When comparing different groups or treatments, the contracting and relaxing responses of biphasic concentration-response curves can be analyzed separately. The maximal relaxation and contraction responses in each phase can be calculated and expressed as a percentage of previously obtained tone ( 474 ). Alternatively, biphasic concentration-responses curves to ACh can be statistically evaluated by ANOVA followed by a post-hoc test to identify possible differences between groups at each ACh concentration ( 475 ). To evaluate the endothelium-dependent contraction to ACh only, vessels with intact endothelium can be incubated with L-NAME and/or EDH inhibitors [please see Table 15 , and section titled “ Endothelium-dependent hyperpolarization (EDH) ”] to prevent the biphasic response and optimize endothelium-dependent contractions ( 476 ). By eliminating the relaxation phase, standard pharmacological parameters (e.g., EC 50 and E max ) can be determined based solely on the concentration–response contraction curve. Pharmacological and Physiological Considerations on Sex Differences in Endothelial Function Sex is a crucial biological variable influencing vascular function and experimental outcomes. Understanding these differences is essential for designing accurate pharmacological and physiological studies. Sex differences in EC biology may reflect inherent chromosomal differences and/or lifelong exposure to sex steroids, as well as gender-related influences in humans ( 477 ). Sex Differences in Vascular Function: Males and females exhibit differences in endothelial function, which affect pharmacological responses to vasodilators and vasoconstrictors. Accordingly, conduit and resistance arteries from adult healthy females exhibit a reduced maximal response (E max ) and a lower EC 50 to different vasoconstrictor agonists when compared with adult healthy males ( Fig. 22 ). The concentration required to induce vasoconstriction (preconstriction) prior to performing relaxation curves may differ between males and females. This is also valid to understanding vasoconstriction. It is recommended to start concentration-response curves at very low concentrations (10 −10 M) when comparing male and female arteries. Thoracic aorta and mesenteric resistance arteries from female animals show an increased ACh-induced relaxation, likely due to enhanced NO bioavailability and the influence of the sex hormones ( 478 )( 479 ). In contrast, bradykinin-induced relaxation does not differ between male and female porcine coronary arteries preconstricted with U46619 ( 480 ). Female rats and mice of the same age are generally smaller than males, but the approach to vessel isolation remains similar, exception for uterine arteries. The uterine artery requires special attention during dissection due to its morphology and physiological variability based on hormonal fluctuations. For further information about uterine arteries, please see ( 1 ). For female animals, it is important to assess the stage of the estrous cycle before using the arteries, as hormonal fluctuations during the cycle can significantly influence vascular function. Although the magnitude of these effects is not universally consistent across vascular beds or vasoconstrictors, the estrous cycle is known to influence sensitivity and NO bioavailability. The estrous cycle comprises proestrus, estrus, metestrus, and diestrus, each defined by distinct estrogen and progesterone profiles ( 481 ): Proestrus / Estrus (high estrogen): enhanced endothelial function, increased NO production, and reduced vasoconstrictor responsiveness. Metestrus / Diestrus (low estrogen): decreased endothelium-dependent relaxation and increased sensitivity to vasoconstrictors. Considerations should also account for human arteries, particularly regarding hormonal status before and after menopause, as well as variations during the menstrual cycle. Clinical evidence parallels these observations, showing menstrual-cycle-dependent variations in endothelial function in women ( 482 ). Ovariectomy markedly alters endothelial function, increasing contraction to angiotensin II and phenylephrine while reducing ACh-mediated relaxation in the aorta and mesenteric arteries ( 483 , 484 ), underscoring the modulatory role of ovarian hormones. Figure 22. Open in a new tab Illustration shows a hypothetical concentration-response curve to a vasoconstrictor in arteries from males (blue) and females (pink). In general, conductance and resistance arteries from healthy adult female mice and rats exhibit a reduced maximal response (E max ) and a lower EC 50 to vasoconstrictor agonists (e.g., noradrenaline and serotonin) compared with age-matched healthy males. Concentration-Response Curves Constructed with Sex Hormones: Estrogen, testosterone, and progesterone induce relaxation in isolated blood vessels at higher concentrations. Generally, relaxation occurs at concentrations greater than 10 μM in vessels pre-contracted with U46619 at approximately 50% of E max . Of note, using such high concentrations of steroid hormones in the absence of binding proteins likely represents a pharmacological condition that may not reflect physiological conditions. The hormone concentration-response curves are typically limited, using no more than six points ranging from 1 μM to 100 μM. U46619-induced contraction provides a stable and sustained vasoconstriction over an extended period (with longer intervals between each addition of a higher hormone concentration), making it the preferred choice for constructing hormone concentration-response curves. Importantly, testosterone has been shown to cause rapid, endothelium-independent vasodilation in several types of blood vessels. Therefore, it is important to use endothelial-derived factor inhibitors or to remove the endothelium to better understand the endothelial contribution to the effects of these hormones (Please see Table 15 ). Overall, when conducting vascular function studies, sex-specific differences must be carefully considered in experimental design. Pharmacological studies should account for differences in vasoconstrictor and vasodilator responses to avoid misleading interpretations. This also applies to physiological approaches, such as pressure myography when performing myogenic response curves, and examining flow-induced dilation, as described above. Sex Differences in Cultured ECs: In addition to whole-artery preparations, sex differences are also evident in isolated ECs, influencing mechanistic, pharmacological, and imaging-based assays ( 485 - 487 ). These differences stem from intrinsic genomic, hormonal, and metabolic factors that persist even when cells are cultured under standardized conditions. Primary ECs isolated from males and females maintain sex-specific transcriptional signatures, including genes involved in inflammation, oxidative stress, angiogenesis, and NO production. Female-derived ECs typically exhibit higher basal eNOS expression and NO bioavailability ( 486 ), whereas male-derived ECs show relatively increased oxidative stress markers. These intrinsic differences persist across early passages and can influence experimental outcomes such as migration, proliferation, and wound-healing assays. Even though cultured ECs are removed from the in vivo hormonal environment, they retain hormone receptor expression (e.g., estrogen receptors α and β, GPER; androgen receptor). Thus, experiments evaluating responses to estrogen, progesterone, or testosterone require careful control of serum content. Charcoal-stripped serum is applied in hormone-sensitive cell culture to decrease endogenous hormones, thus guaranteeing that observed effects are uniquely attributable to exogenous-added hormones or compounds. Importantly, estrogen exposure can acutely increase NO production and modulate Ca 2+ dynamics, whereas testosterone often triggers rapid, largely endothelium-independent vasodilation mechanisms that can also be observed in cultured ECs. Sex-based differences extend to intracellular Ca 2+ handling ( 485 ). Accordingly, in female ECs, transient receptor potential vanilloid 4 and mitochondrial Ca 2+ uniporter increase Ca 2+ -dependent NO production. In males, only transient receptor potential canonical 3 plays a fundamental role in this effect. Further, it was also shown that mitochondrial function in ECs differs by sex, where female mice having enhanced Ca 2+ uptake capacity, and that these differences are attributable to the presence of more mitochondria and a higher mitochondrial membrane potential in female mice rather than differences in composition of the mitochondrial Ca 2+ uniporter complex. Therefore, when performing Ca 2+ imaging experiments (e.g., with Fura-2, Fluo-4, GCaMP reporters), it is essential to analyze male and female ECs separately. Agonists such as ACh, ATP, histamine, and shear stress mimetics may produce different Ca 2+ kinetics across sexes. Factors such as culture duration, passage number, and confluency can interact with sex-specific Ca 2+ control mechanisms and should be standardized. Assays such as tube formation, migration, barrier integrity, oxidative stress measurements (MitoSOX, DHE), and mechanical stimulation (e.g., shear stress) may all yield sex-dependent variations. Accordingly, it has been shown that female HUVEC has a higher cell viability after serum starvation and an increased tube formation capacity compared to male cells ( 488 ). Regarding ECs permeability, it was observed that estrogen treatment protects the brain from the immune response under inflammatory conditions by enhancing BBB (formed primarily of brain microvascular ECs) functionality and decreasing leukocyte extravasation across the BBB ( 489 ). Overall, sex is a critical biological variable in cultured EC studies, as male- and female-derived ECs retain intrinsic genomic, metabolic, and hormonal differences that directly influence experimental outcomes. These sex-specific properties persist despite standardized culture conditions and affect a wide range of assays, including Ca 2+ imaging, migration, proliferation, tube formation, oxidative stress, and barrier function. Therefore, experimental design should incorporate sex as a factor, analyze male and female ECs separately, and standardize variables such as passage number, serum composition, and confluency to avoid confounding effects. Recognizing and accounting for these persistent sex-based differences is essential for generating reproducible, physiologically relevant, and accurately interpretable EC data. 4. SOFTWARE ACQUISITION & ANALYSIS Several software platforms are available for analyzing data acquired in EC function studies. Traditionally, these have been proprietary software solutions that are bundled with or sold as an add-on to commercial acquisition software and equipment. However, scientists have long developed their own tools, often sharing them freely within research communities. In fact, many of the currently-available commercial programs used in endothelial studies originated in research laboratories and continue to be developed in collaboration with researchers. The growing momentum of Open Science principles has expanded this collaborative practice beyond the sharing of working software to include the publication and sharing of open-source, documented code and tools that can be freely used and adapted. This practice encourages greater transparency, reproducibility, and collaborative improvement of analytical tools. The result is that researchers studying endothelial function today have access to a diverse ecosystem of freely distributed or fully open-source software tools that can be adapted, extended, improved, and even reinvented by the broader scientific community. In the following sections, we present an overview of the main software options available for endothelial function research, including commercial solutions, freely distributed tools, and open-source software. Importantly, across endothelial function assays, including pressure myography, wire myography, pin myography, and related approaches, modern instrumentation typically provides digital, automated data acquisition (e.g., digital edge-detection for diameter or digital force transducers for tension), which minimizes potential bias and/or influence. However, some laboratories still use analog or manual output methods, such as manual diameter tracking, video calipers, or chart-based tension recordings. In these cases, because data acquisition involves greater subjectivity, measurements should be performed by a blinded investigator to reduce potential bias when comparing experimental groups. Software for Pin & Wire Myography In the pin and wire myographs, the primary measurement of interest is isometric tension (force) generated by VSMCs in response to stimuli. This tension increases during VSMC contraction, decreases during relaxation, and is measured using force transducers. For these organ bath pharmacology experiments, software must convert the analog transducer signal to digital data in appropriate units of force (e.g. grams or milli-Newtons) while also enabling real-time visualization and analysis of the data to allow researchers to monitor and annotate their experiments as they progress. In addition to force transducers, other physiological parameters such as temperature may be monitored, and software may also provide control features, such as electrical stimulation. But at their most basic, all pin/wire myography software must provide a live readout of force in a user-friendly graphical display. For experiments with a single force transducer/myograph, the freely-distributed Chart Software by Strathclyde Electrophysiology Software (SES) does this. However, wire myographs are available in multi-channel versions allowing researchers to perform parallel experiments in up to four different blood vessels using a single integrated system. For these systems, commercial software packages provide the additional functionality of measuring from multiple force transducer channels simultaneously. The most common commercial software packages for multi-channel recording are: LabChart: a flexible software from ADInstruments. LabScribe: from iWorx. MyoDAQ: proprietary software from Danish Myo Technology. Each of these packages facilitates easy exporting of key data directly to graphing software for plotting and further analysis. Several of them also offer additional analysis add-ons, such as peak analysis, and concentration-response modules to enhance data interpretation and experimental workflow. Software for Pressure Myography In the pressure myograph, the primary measurement of interest is the inner or outer diameter of blood vessels visualized via light microscopy. The success of these experiments relies on software that can capture and analyse images in real time to track vascular responses accurately. Unlike simple measurement tools, pressure myograph software must process video streams, apply automated diameter tracking analysis, and allow seamless integration with other physiological parameters (e.g. temperature and pressure). The core of pressure myograph software is the ability to process video streams of blood vessel images and apply automated edge detection algorithms to continuously track vessel diameter throughout the duration of an experiment. The software must also display this diameter information graphically, allow the user to annotate the experimental data trace to indicate when pharmacological manipulations were performed, and provide feedback to the user on the state of the artery at any moment in time, such as displaying current percentage diameter relative to control conditions. Pressure myograph software is available from several laboratories ( 490 )( 491 )( 173 ). One of these, VasoTracker, is an open-source software specifically designed for real-time blood vessel diameter tracking. VasoTracker was first released as part of a fully open-source pressure myography system, providing researchers with a cost-effective, flexible and customizable alternative to proprietary tools, allowing them the freedom to build and modify their own myograph setups. The latest software release, VasoTracker 2 ( 492 ) ( Fig. 23 ) is now fully integrated with μManager microscope control software ( 493 ) allowing diameter tracking software to be added to existing microscope systems, including pressure myographs from commercial suppliers. The software supports both live diameter tracking and offline analysis of pre-recorded images, making it highly versatile. Researchers can define multiple regions of interest, user-defined scan lines, and customize various settings, ensuring compatibility with a range of experimental designs and imaging modalities, including fluorescence and ultrasound imaging. Figure. 23. Open in a new tab Screenshot of the VasoTracker 2 myography and blood vessel diameter measurement software. The image shows a trace generated by analysing a pre-recorded video of a flat-mounted rat mesenteric artery, where endothelial cells were loaded with a fluorescent indicator. The trace represents changes in vascular tone during contraction to phenylephrine and subsequent relaxation to acetylcholine. VasoTracker 2 can analyse vessel diameter changes from both live imaging and pre-recorded videos. The most widely used commercial software packages for pressure myography are: MyoVIEW: proprietary pressure myograph software from Danish Myo Technology IonWizard: advanced multi-functional software from IonOptix with a diameter measurement addon (VesAcq) These software packages are generally sold with complete myograph systems and provide much of the same functionality as the open-source/academic software available. Software for Fluorescence Imaging Many aspects of assessing EC function rely on the use of fluorescence indicators visualized via microscopy. For example, redox-sensitive probes can be used to assess oxidative stress ( 140 ), NO indicators provide a readout of vasodilator production ( 494 ), and membrane potential probes provide a readout of electrical signaling ( 495 ). Perhaps the most widely used functional assay of endothelial function is Ca 2+ -sensitive dyes. Please refer to Section G, “EC Ca 2+ Imaging” , and Table 6 for further details on Ca 2+ -sensitive dyes. These indicators allow researchers to monitor Ca 2+ levels in response to vasoactive stimuli. In endothelial research, all of these fluorescence-based tools have been used in various preparations, including cultured ECs, freshly isolated ECs, and intact arteries, whether pressurized and visualized via confocal microscopy or in the flat-mount en face or inverted artery preparation. This versatility makes fluorescence-based imaging a cornerstone technique in endothelial function studies. The acquisition software used by endothelial researchers varies wildly, typically depending on the specific microscope system used. Common proprietary software includes: NIS-Elements for Nikon microscopes AxioVision for Zeiss FluoView for Olympus. Academic software is also available for fluorescence imaging. For example, the freely distributed SES WinFluor Software ( 496 ) is designed for simultaneous fluorescence imaging and electrophysiological recordings. But whilst software like WinFluor provides specialized functionality, the dominant microscope control and imaging software is now μManager ( Fig. 24 ). Figure 24. Open in a new tab μManager microscope control software and ImageJ/FIJI analysis. The screenshot illustrates the process of image acquisition and analysis using μManager and ImageJ (or FIJI). On the left, μManager 2.0 is used to record fluorescence time-series of an isolated patch of endothelial cells loaded with the fluorescence superoxide indicator, dihydroethidium. On the right, a previous recording has been analysed using built in ImageJ tools to plot fluorescence intensity over time. In this experiment, the increase in dihydroethidium fluorescence was caused by the mitochondrial uncoupler, CCCP. μManager is an NIH-funded, open-source platform that is built on ImageJ ( 493 )( 497 ), providing seamless integration with its analysis algorithms and plugins. This allows researchers to control microscopes, acquire images, and immediately process them with ImageJ. μManager is supported by nearly all microscope device manufacturers, making it the most widely used alternative to proprietary imaging software. Software for Fluorescence Imaging Analysis Fluorescence-based imaging in endothelial research is used to measure intracellular signals, but the specific analysis required depends on the experimental approach. The most basic measurement often involves comparing fluorescence intensity before and after an experimental treatment, or between control and diseased tissue. For experiments involving dynamic signaling processes, such as Ca 2+ imaging, fluorescence intensity may be continuously recorded to track real time changes and allow visualization of temporal signals rather than just endpoint comparisons. Particularly in Ca 2+ imaging, researchers may be interested in assessing the signals arising from individual cells, or even in subcellular analysis of localized signaling dynamics within individual ECs. Regardless of the specific experimental probe, the most widely used image analysis software for assessing fluorescence images of ECs is ImageJ and the biological-image analysis focused distribution, FIJI ( 498 ). ImageJ/FIJI provides a comprehensive suite of tools for fluorescence image analysis, making it suitable for any experimental design. It allows researchers to quantify fluorescence in individual cells or across tissue samples. Users can apply background correction and noise reduction filters to enhance signal detection, segment and track individual cells or regions of interest (ROI) using the ROI tool, and even analyze time-series data for dynamic imaging experiments. For additional guidance on best practices and available analysis tools, readers are encouraged to consult the official ImageJ/FIJI documentation, which provides detailed recommendations for Ca 2+ imaging workflows. Software for Endothelial Ca 2+ Imaging Analysis Although ImageJ/FIJI remains the gold standard for fluorescence-based imaging, it is rarely used for Ca 2+ imaging analysis due to the manual nature of its tools, which can be time-consuming and prone to user bias. To address this, researchers have typically developed automated solutions tailored to specific experimental needs. These range from studies of population-wide Ca 2+ dynamics, to whole-cell signals, and to subcellular, elementary Ca 2+ events, with both fully open-source and commercial tools widely used for EC research. Several open-source analysis tools have been developed specifically for analysis of cell-wide endothelial Ca 2+ signaling. LC_Pro is an ImageJ plugin designed for whole-cell Ca 2+ event analysis in ECs ( 499 ). Similarly, the S8 algorithm from the same authors is implemented in Python ( 499 ). Both automatically generate ROIs for each detected Ca 2+ event, making them particularly useful for unbiased event detection, but are incapable of providing information regarding the percentage of cells activated. Another algorithm is WAVE (whole-cell average), a Python implementation developed specifically for analyzing cell-wide and networked analysis of Ca 2+ responses in large-population EC imaging ( 500 )( 501 ). It enables longitudinal tracking, allowing responses to be matched across cells over time and imaging sessions, but has not yet been released as a fully open-source tool. At the subcellular level, Ca 2+ signaling events include Ca 2+ sparks (ryanodine receptor-mediated), puffs (IP 3 receptor-mediated events) and sparklets (influx-mediated events). SparkMaster ( 502 ), developed as an ImageJ plugin, was one of the first automated tools for elementary Ca 2+ event detection, designed to analyze Ca 2+ sparks imaged by confocal line-scan microscopy. This has been followed by the recent SparkMaster 2 ( 503 ), a python-based update offering improved automation and analysis features. SparkAN is an in-house package designed for similar analysis of elementary IP 3 -mediated Ca 2+ release and Ca 2+ influx events, but from three-dimensional (XYT) confocal imaging ( 104 )( 122 ). Similarly, FLIKA is a Python package designed for analyzing elementary IP 3 -mediated Ca 2+ release events in TIRF imaging ( 504 ) that works equally well for high-resolution epifluorescence imaging of ECs ( 505 ). Despite these developments, there is not a widely available, user-friendly & open-source platform for endothelial Ca 2+ imaging analysis. Tools like Caiman ( 506 ), offer advanced computational approaches, but a more accessible solution, potentially implemented in Napari ( 507 ), could provide interactive, scalable, and easy-to-use analysis for EC Ca 2+ signaling. Software Beyond Myography and Fluorescence Imaging Beyond myography and fluorescence imaging, computational tools are essential for analyzing endothelial function at multiple levels. For example, flow cytometry and transcriptomic profiling are widely used to define endothelial phenotypes, with open-source tools such as FlowJo (for cytometry), Scanpy (Python) and Seurat (R) enabling large-scale data analysis. These open-source computational approaches are increasingly important, not only for handling complex datasets, but also for ensuring transparency, shareability, and reproducibility in endothelial research. At the same time, much of endothelial research relies on routine data analysis from imaging, functional assays, and biochemical experiments, where tools such as Excel, Origin, and GraphPad Prism remain widely used for visualization and statistical analysis. Making specialized computational pipelines and more routine data analysis more accessible and standardized will help improve reproducibility across EC research. X. FUTURE DIRECTIONS & EMERGING TECHNOLOGIES Potential areas for future research and development Challenges for Induced Pluripotent Stem Cells-Derived Induced EC: Lineage Subtypes, Tissue Specificity, and Functional Validation Since induced pluripotent stem cells (iPSCs) were generated by Yamanaka et al. using four transcriptional factors (e.g., OCT4, SOX2, KLF4, and C-MYC, or OSKM factors) ( 508 , 509 ), iPSC-induced ECs (iECs) have emerged as a source of patient-specific or disease-relevant ECs. Compared with embryonic stem cell-derived iECs, iPSC-derived iECs confer several advantages including free from ethical concerns and using autologous sources. In this section, we will mainly discuss iPSC-derived iECs and their potential applications as novel preclinical models for mechanistic and drug screening studies of vascular functions. The general differentiation strategy of iPSC to iECs is to culture iPSCs on a Geltrex-coated surface and various growth factors and chemicals will be added in a timed fashion to progressively drive the differentiation course followed by molecular and functional validations ( 510 ) ( Fig. 25 ). Three differentiation phases are comprised: primitive streak mesoderm, lateral mesoderm/progenitor cells, and mature iECs. BMP4, FGF2, and ACTIVIN A are the common factors in inducing early mesoderm, followed by continuation of BMP4, FGF2, and VEGF for driving endothelial progenitor cells (EPCs). GSK-3 inhibitors such as CHIR9901 are commonly used to stimulate the mesoderm differentiation by promoting canonical Wnt signaling ( 510 )( 511 ) ( 512 ) ( 513 ). The purification and selection of EPCs using KDR, CD31, and CD34 biomarkers can be used to enrich EPC populations, thus increasing the differentiation efficiency of iEC lineages ( 513 )( 514 )( 515 )(572). In the final step, iECs will be induced using FGF2, VEGF, and a TGFβ inhibitor SB431542. Inhibition of TGF-β signaling is crucial to drive mesoderm cells to endothelial specification, suppress endothelial-to-mesenchymal transition, and maintain EC identities ( 512 ). The course takes about 10 to 14 days. Studies have shown that many factors can improve the differentiation efficiency of iPSC to iECs, including MEK/ERK inhibitor, the γ-secretase inhibitor DAPT, miR-495, RNA binding protein Quaking isoform 5, and cAMP ( 512 )( 515 )( 516 )( 517 )( 518 )( 519 )( 520 )( 521 )( 522 )( 523 )( 524 )( 525 )( 526 ). However, discrepancy has been reported with suppression of iEC differentiation by MEK/ERK inhibitor PD0325901 ( 527 ). Other factors such as ECM compositions and shear force can regulate iEC differentiation ( 518 )( 519 ). Figure 25. Open in a new tab Schematic of capillary malformation (CM) iPSC generation, iEC differentiation and functional validation. CM skin lesions were used for outgrowth of human dermal fibroblasts, followed by reprogramming into iPSCs. The stem cell biomarkers Nanog and Tra1-60 were used to verify the molecular features of iPSC colonies. CM iECs were differentiated and characterized by pan EC biomarkers, CD31 and CD144. Further functional validation of iECs was performed using in vitro tube formation assay and in vivo xenograft plug-in assay. Image taken with permission from ( 510 ) There are two main challenges for iEC differentiation. First, whether iECs can be induced to specific lineage commitment, which will depend on environmental cues such as fluid flow, pulsatile pressure, and compositions of ECM that are insufficiently incorporated in the differentiation course of iPSCs to iECs. Rufaihah et al. ( 520 ) found that iECs displayed intermediate phenotypes and functional heterogeneity with arterial, venous, and to a lesser degree, lymphatic lineage biomarkers. To generate lineage specific iECs, types and doses of VEGF and other co-factors were optimized: a lower concentration of VEGF-A favored venous lineage, a higher concentration of VEGF-A and 8-bromoadenosine-3':5'-cyclic monophosphate promoted arterial subtype, whereas VEGF-C and angiopoietin-1 facilitated lymphatic EC development ( 520 ). Zhang et al. induced arterial iECs using a “five factor” protocol ( 521 ): 100 ng/mL FGF 50 ng/mL VEGF 10 μM SB431542 5 μM RESV 10 μM L690). Sivarapatna et al. used a biomimetic flow bioreactor to mimic shear force for inducing arterial iECs ( 522 ). Ang et al. induced arterial iECs by inhibiting vein-specifying signals (e.g., PI3K) and vice versa and resulted in >90% pure human arterial or venous iECs from iPSCs ( 523 ). They found a temporally-dynamic role for TGFβ signaling in specifying arterial fate. TGFβ needed to be initially activated for inducing primitive mesoderm streak, then inhibited for specifying lateral mesoderm, and later re-activated to induce arterial fate ( 523 ). Lee et al. generated lymphatic iECs using VEGF-A, VEGF-C, and EGF. The lymphatic iECs expressed LYVE-1 and PODOPLANIN and could form lymphatic vessels in vivo ( 524 ). Second, whether iECs can be differentiated into tissue-specific subtypes. One main feature of ECs is their tissue heterogeneity, leading to distinct tissue-specific functions and responses. Many efforts have been carried out to examine whether iECs can display certain tissue-specific phenotypes under tailored differentiation conditions. Linville et al. ( 525 ) differentiated iPSCs into a mixed EC/neural progenitor cell under unconditioned media without bFGF and further generated brain microvascular iECs with all-trans retinoic acid in the induction medium. The induced brain iECs were seeded within templated type I collagen channels and shear stress was applied for formation of microvascular network. These iECs expressed BBB-specific tight junction proteins, including Claudin-5, Occludin, and ZO-1, the transporter GLUT-1, and the efflux transporter P-glycoprotein. Stebbins et al. ( 526 ) used the same strategy and recapitulated the induced brain microvascular iECs. Another example is to induce corneal iECs from iPSCs. Neural crest cells were first differentiated from iPSCs, followed by corneal iEC induction with WNT activation, TGF-β inhibition, and addition of retinoic acid. The iPSC-derived corneal iECs expressed corneal EC-specific biomarkers such as ATP1A1 and ZO-1 and exhibited hexagonal/polygonal morphology ( 528 ). The authors also found that coating with vitronectin could increase the efficiency of corneal iEC differentiation ( 526 ). In a separate study, choroidal iECs were differentiated with the addition of connective tissue growth factor toward a choroidal EC fate ( 529 ). Prasain et al. developed a protocol to sort NRP-1 + CD31 + cells from mesoderm cells and drive them into cord-blood endothelial colony–forming cells (ECFCs). These cells showed high clonal proliferative potential and the capacity to form human vessels in vivo ( 530 ). Procedures for iEC Validation The major challenge for clinical applications of iECs is lack of a standard for quality evaluation of iECs. The iECs generated from iPSCs through various differentiation protocols display many common EC phenotypes, such as expression of pan EC biomarkers of CD31, KDR, and CDH5, formation of in vitro 2-D capillary-like structures on Matrigel, uptake of LDL, and proliferation in vitro. However, there are many factors hampering potential applications of iECs as future therapeutics, including different cell sources used for iPSC generation, vast numbers of variable differentiation protocols from different laboratories, potential heterogeneity of iECs generated under distinct protocols, and necessity of diverse cell types and environmental cues for ECs to achieve similar maturation and functionality as in vivo . These concerns and shortcomings have raised the need for a guideline for functional and phenotypic validations of iECs for laboratory practice. Here, we attempt to propose a general guideline for quality controls of iECs. The following assays can be considered for validation of general function and phenotype of iECs: a. Including a control of commercially available primary human ECs from the same tissue as the source of iPSCs ( 531 )( 532 ). b. Using immunofluorescent staining or flow cytometry to determine expressions of pan EC biomarkers (CD31, CDH5, eNOS, KDR, vWF) ( 510 )( 520 )( 532 )( 533 ) c. Assessing proliferative and migrative rates of iECs ( 533 ). d. Using in vitro tube-formation assay to evaluate the capability of iECs to assemble capillary-like structures on Matrigel or Hydrogel with morphological measurements of diameter, thickness of branches, and branch points ( 513 )( 532 )( 534 ) e. Capability of LDL uptake ( 531 )( 535 ). Please see section L. “Approaches to Assess Endothelial Metabolism” . f. Endothelial NO synthesis using fluorescent dyes such as DAF-2 DA and DAF-FM DA for in situ NO detection ( 531 ). g. Evaluating endothelial barrier function using transendothelial electrical resistance or permeability assays ( 533 ). Please see section H. “Endothelial Barrier Integrity” . h. Examining junctional integrity upon shear stress stimulation by detecting adherence and tight junction components ( 536 ) i. Assessing glycolytic metabolism of iECs including rates of glycolysis, glycolytic capacity, and glycolytic reserve ( 531 ). Please see section L. “Approaches to Assess Endothelial Metabolism” . j. Determining iEC capability of in vivo vessel formation ( 510 ). Xenograft implantation is the common in vivo assay by injecting iECs subcutaneously or into various disease models such as ischemia ( 537 )( 538 ), myocardial infarction ( 517 ), wound healing ( 524 ), and retinopathy ( 528 )( 530 ). The functional incorporations of iECs can be assessed at different time points post-implantation. This method will assess if donor iEC-derived vessels can incorporate into the host vasculatures to exert beneficial effects in these models. The animal strain, sex, age, and genetic backgrounds are variables to be considered. Depending on the lineage and tissue specificity, additional functional and phenotypic validation assays for lineage- or tissue-specific iECs can be included: (1) expression of specific venous (NR2F2, EPHB4), arterial (NOTCH1, DLL4, CXCR4, EFNB2, HEY2, ANTXR1) or lymphatic (LYVE-1, PROX1, PDPN) biomarkers ( 520 ); (2) tissue specific EC biomarkers such as brain microvascular EC biomarkers Claudin-5, Occludin, ZO-1, and GLUT-1 ( 525 ); (3) surface immune biomarkers of ICAM-1, ICAM-2, and E-selection and monocyte adhesion endothelial disturbed flow assay ( 539 ); Many disease-sourced iPSC-derived iECs recapitulate pro-inflammatory signaling ( 510 )( 540 ) from original lesion cells, thus interactions between iECs and immune cells are critical for determining pathogenic factors inherited by iECs; (4) EC colony formation assay for the proliferative potential ( 530 ). Like mature primary ECs with restricted capability of dividing, monolayer iECs generated from various protocols have exhibited limited proliferative potential (up to 5-8 passages to reach senescence or undergo EndMT). A high proliferative potential ECFC generated from iPSCs may include many progenitor cells that can highly proliferate and commit to iECs; and (5) multiome studies including RNA-seq, ATAC-seq, and ChIP-seq to reveal the molecular phenotypes and enriched pathways for iECs ( 510 )( 533 ). Overall, iPSC-derived iECs have emerged as novel resources for their potential in regenerative medicine. Particularly, many disease-specific iPSCs and their iECs have been established and characterized. These cells have become clinically relevant models for disease pathogenesis exploration and drug discovery. A more specific guideline for iEC molecular and functional validation will be necessary for using these valuable models studying the detrimental factors to EC dysfunction under disease conditions and subsequent drug screening. Incorporation of Novel Technologies in Evaluating Endothelial Function The assessment of endothelial function has evolved significantly with the advent of innovative technologies such as microfluidics and organ-on-a-chip (MOOC) models, advanced imaging modalities, and multi-omics approaches. These tools offer unprecedented opportunities to study endothelial biology at higher resolution and throughput, and in more physiologically relevant contexts. This section outlines key considerations for integrating these novel technologies into experimental evaluations of vascular endothelial function. Microfluidics, Organ-On-A-Chip models, and Endothelialized Whole Organ Scaffolds Two-dimensional in vitro models usually lack the complex vascular geometries and cell heterogeneity seen in vivo . MOOC devices offer clear advantages, such as providing a complex geometry with the inclusion of fluid flow, which is a physiologically relevant model for studying endothelial responses in disease progression ( 541 ). For example, extracellular matrix can be mimicked in MOOC devices by adding the naturally-derived proteins or synthesized biomaterials including fibronectin, collagen with different stiffnesses to study the vascular stiffness ( 541 )( 542 ). Leukocytes, platelets and inflammatory cytokines can be perfused into the MOOC devices coated with vascular ECs to investigate the leukocytes/platelets and EC interactions under the inflammatory environment in the process of atherosclerosis and sepsis development ( 543 )( 544 ) Moreover, MOOC devices can also be used to assess endothelial barrier integrity, permeability, and responses to mechanical stimuli or pharmacological agents ( 541 ). These platforms are particularly valuable for real-time monitoring of EC behavior under controlled flow conditions. Traditional culture and device systems do not adequately mimic the native microenvironment and often fall short in accurately modeling human pathophysiology for studying diseases and drug mechanisms. Whole-organ decellularization offers a strategy to create a scaffold that preserves the structural, mechanical, and biological characteristics of an intact vascular network ( 545 ). Yuan et al. optimized a culture protocol to improve EC coverage in decellularized whole lung scaffolds within a biomimetic bioreactor ( 546 ). Single-cell RNA sequencing revealed that primary pulmonary microvascular ECs partially regain native heterogeneous endothelial phenotypes including general capillary and aerocyte ECs. Similarly, human iPSC-derived ECs cultured in this system began exhibiting native human endothelial traits, restoring vascular barrier function and maintaining small capillary patency. To assess functional responses, LPS treatment was applied to simulate acute lung injury, leading to increased proinflammatory signaling and vascular barrier impairment, demonstrating the system’s ability to model inflammatory endothelial dysfunction. Advanced Imaging Modalities Live and deep histological imaging techniques including multiphoton/intravital microscopy, optical coherence tomography (OCT), and fluorescence-based imaging provide detailed insights into endothelial structure and dynamics. Advanced imaging modalities have significant implications for diagnosing vascular diseases, where endothelial dysfunction often serves as an early hallmark. OCT can detect endothelial injury, plaque buildup, and vessel wall thickness, aiding in the diagnosis of atherosclerosis and guiding stent placement ( 547 )( 548 ). Similarly, fluorescence-based imaging, often coupled with targeted probes (e.g., VEGFR antibodies labeled with fluorophores), enables the detection of molecular changes in ECs associated with diseases like tumor vasculature ( 549 ). Nuclear imaging techniques, such as positron emission tomography (PET), provide functional insights by tracking metabolic activity or inflammation within the vasculature. For example, 18 F-FDG PET-MRI has been used to identify active atherosclerotic plaques by detecting endothelial inflammation ( 550 )( 551 ). These diagnostic capabilities underscore the translational potential of advanced imaging, moving beyond research to inform precision medicine in vascular pathologies. A standout feature of intravital multiphoton microscopy is its ability to dissect complex cellular interactions within the vasculature. Schilling et al. demonstrate the dynamic changes of vessel types (arterial, venous, and capillary vessel) networks at the site of cranial bone defect repair via multiphoton microscopy ( 552 ). Using advanced longitudinal intravital multiphoton microscopy, Bixel et al. demonstrate that early vascular sprouting during calvarial bone regeneration is not directly coupled to osteoprogenitor invasion. Instead, osteoprogenitors originating from the periosteum differentiate into bone-forming osteoblasts at the injury site ( 271 ). While live imaging excels at dynamic processes, techniques like tissue clearing combined with light sheet microscopy have transformed our ability to map vascular networks at a whole-body or organ scale ( 553 )( 554 ). This combination has been used to map the entire vascular network of mouse heart under myocardial infarction and reveal endothelial plasticity which drives aberrant vascularization ( 555 ). For example, studies employing light sheet microscopy with cleared mouse brains have been used to determine detailed alterations of cerebrovascular networks in aged mice. Light-sheet imaging combined with 3D immunolabeling reveals greater arteriole tortuosity in aging brains. Notably, vascular and pericyte densities exhibit significant and selective declines in the deep cortical layers, hippocampal circuitry, and basal forebrain regions ( 556 ). Multi-Omics Approaches Single-cell transcriptomics have been widely used to identify the endothelial heterogeneity in the past few years. Single-cell EC transcriptomes from 11 mouse tissues were generated with 78 EC subclusters being identified. ECs from various vascular beds (arteries, capillaries, veins, and lymphatics) displayed transcriptomic similarities across tissues, while EC heterogeneity was primarily influenced by tissue type rather than vessel type ( 557 ). For further information about EC heterogeneity, please see section above “ The Heterogeneity of Endothelial Cells ”. Single-cell transcriptomics require tissue dissociation, leading to a biased selection of captured cells and the loss of morphology and spatial context. Spatial transcriptomics are increasingly integrated with single-cell transcriptomics and tissue imaging to map the molecular profile within spatial context. Two primary spatial transcriptomics methods have been developed for detecting mRNA transcripts using different strategies: multiplexed fluorescent in situ hybridization (FISH) techniques such as MERFISH and Xenium and in situ capture followed by high-throughput sequencing such as Visium and Stereo-Seq ( 558 ). For example, Liu et al. ( 559 ) identified that lung general capillary ECs (gCap) in the microvascular bed can transdifferentiate into arterial ECs, contributing to the development of pulmonary arterial hypertension. The multi-omics data from genomics, transcriptomics, proteomics, and metabolomics can be effectively integrated using robust algorithms such as pathway enrichment analysis, machine learning, or network modeling to extract meaningful insights. For instance, multi-omics profiles of patient-derived samples could help predict the repurposing of FDA-approved medications using the Connectivity Map dataset ( 560 ). Additionally, self-supervised and unsupervised learning techniques, including large language models and deep learning for image analysis, may facilitate the unbiased discovery of biological processes ( 554 ). XI. CONCLUSION The assessment of endothelial function remains a cornerstone in understanding vascular physiology and pathology. Through the collective efforts of leading investigators across multiple countries, these guidelines were developed to establish best practices for evaluating EC function in vascular tissue. We integrated traditional approaches, such as wire and pressure myography, with emerging molecular and imaging methodologies that reflect the complexity and dynamic nature of EC biology. While we made a concerted effort to include a diverse group of vascular biologists with at least 15 years of experience in endothelial research, we acknowledge that numerous outstanding contributions could not be featured due to scope limitations. We strongly encourage readers to explore additional studies and innovative methodologies beyond those included here to further enrich their understanding. Importantly, while the guidelines reflect a broad consensus on most techniques, a few differences remain due to varying laboratory traditions, for example, the choice of solutions or terminology. Nevertheless, we ensured that only proven, successful factors were included, with flexibility for adaptation as needed. We aimed to use solutions that are as physiological as possible, as shown in Table 2 ; however, in a few instances, experimental requirements or established laboratory traditions necessitated modifications, which may still differ across settings. These guidelines serve as a resource not only for experienced investigators but also for individuals establishing new laboratories and for students and trainees eager to learn about the tools and techniques critical to studying endothelial function. We recommend that those entering the field embrace both conventional and novel approaches, maintain rigorous methodological standards, and remain attentive to the biological heterogeneity and evolving landscape of endothelial science. By fostering a culture of rigor, transparency, and innovation, we hope these guidelines will help shape the next generation of vascular biologists and promote continued discovery in endothelial research. As you will notice, we also updated the definition of endothelial dysfunction to reflect new discoveries, including the recognition of endothelial phenotypic changes and other dynamic aspects of endothelial biology. Unfortunately, endothelial dysfunction remains a central hallmark of virtually all cardiovascular diseases. We hope that these guidelines not only assist scientists in advancing their research but also help them tackle the formidable challenge of reversing endothelial dysfunction, a crucial step toward treating and, perhaps one day, curing many cardiovascular diseases. Figure 7. Open in a new tab Endothelial Tube Preparation. Endothelial tube segments visualized under a microscope to confirm removal of smooth muscle cells. Morphology of endothelial tubes maintained at (A) 24 °C, (B) 32 °C, and (C) 37 °C. At 37 °C, individual cells retracted away from each other with multiple rounded (i.e., contracted) cells present. Areas that appear out of the focal plane identify gaps that have opened in the upper layer of the collapsed tube. Scale bars = 50 μm. Image taken with permission from ( 125 ). ACKNOWLEDGEMENTS We thank Dr. Alfredo Sanchez Solano, University of Rochester Medical Center, for providing valuable technical feedback on patch-clamp electrophysiology in endothelial cells. We would like to thank Dr. Luciana Venturini Rossoni, Department of Physiology at the Institute of Biomedical Sciences, University of São Paulo, Sao Paulo, Brazil, for her detailed revision and edits to the guidelines. Some of the illustrations were created with Biorender.com. FUNDING NIH (NHLBI R01HL149762) and (NIA R21AG085331-01), Alzheimer’s Association (AARG-NTF-23–1145090) to CFW NIH (NHLBI R01HL168568) to JACG NIH (NHLBI R00HL143198 and R01HL175083) to SDZ NIH (NHLBI R01HL160752) to JKF NIH (NHLBI R35HL155008) and (NINDS R33NS115132) to SE NIH (S10OD023438 and NHLBI R01HL165124) to AJT NIH (NHLBI 5R37HL041026 and NICHD 1R21HD110771-01A1) to SSS NIH (NINDS R01NS134690 and AHA CDA 931652) to CEN NIH (NHLBI R00HL151889, R56HL169223, NIGMS P20GM109091-Pilot Project, P20GM103641-Pilot Project), USC ASPIRE Award (64136) to CGM NIH (NHLBI HL173549 and HL157025) and American Heart Association (AHA9639591 and AHA 863107/847970) to AMB NIH (NIAMS R01AR073172 and 1R21AR083066), Department of Defense (DoD HT9425-23-1-008) to WT NIH (NHLBI R01HL155618) to BEI and PB and NIH (Office of the Director R21OD037867) to PB NIH (NHLBI R01HL158596, R01HL162794 and R01HL170096) to ZD Novo Nordisk Foundation (NNF21OC0071822), Fonden til Lægevidenskabens Fremme (L-2022-00211 and L-2023-00179), and Aarhus University Research Foundation (AUFF-E-2021-9-18) to EB Lundbeck Foundation Grant (R307-2018-3667) and Novo Nordisk Foundation (NNF0095653) to JK Lundbeck Foundation Grant (R323-2018-3674) to TAJ American Heart Association (AHA 916031) to TJC FAPESP (2023/17022-9, 2020/09799-5) and CNPq (fellowship 312126/2022-0) to APD British Heart Foundation (RG/F/20/110007, PG/20/9/34859) to CW Agencia Estatal de Investigación (PID2020-116498RB-I00; MCIN/AEI/10.13039/501100011033) and by ‘ERDF A way of making Europe’; the European Union's Horizon 2020 research and innovation programme under the Marie Sklodowska-Curie grant agreement no. 954798; and Instituto de Salud Carlos III (AC23_2/00044) under the umbrella of the Partnership Fostering a European Research Area for Health (ERA4Health) (GA N° 101095426 of the EU Horizon Europe Research and Innovation Programme) to AMB. Footnotes DISCLOSURES None References 1. Wenceslau CF, McCarthy CG, Earley S, England SK, Filosa JA, Goulopoulou S, Gutterman DD, Isakson BE, Kanagy NL, Martinez-Lemus LA, Sonkusare SK, Thakore P, Trask AJ, Watts SW, Webb RC. Guidelines for the measurement of vascular function and structure in isolated arteries and veins. Am J Physiol Heart Circ Physiol 321: H77–H111, 2021. doi: 10.1152/ajpheart.01021.2020. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 2. Furchgott RF, Zawadzki JV. The obligatory role of endothelial cells in the relaxation of arterial smooth muscle by acetylcholine. Nature 288: 373–6, 1980. doi: 10.1038/288373a0. [ DOI ] [ PubMed ] [ Google Scholar ] 3. Vanhoutte PM. How We Learned to Say NO. Arterioscler Thromb Vasc Biol 29: 1156–60, 2009. doi: 10.1161/ATVBAHA.109.190215. [ DOI ] [ PubMed ] [ Google Scholar ] 4. Vanlandewijck M, He L, Mäe MA, Andrae J, Ando K, Del Gaudio F, Nahar K, Lebouvier T, Laviña B, Gouveia L, Sun Y, Raschperger E, Räsänen M, Zarb Y, Mochizuki N, Keller A, Lendahl U, Betsholtz C. A molecular atlas of cell types and zonation in the brain vasculature. Nature 554: 475–480, 2018. doi: 10.1038/nature25739. [ DOI ] [ PubMed ] [ Google Scholar ] 5. Gillich A, Zhang F, Farmer CG, Travaglini KJ, Tan SY, Gu M, Zhou B, Feinstein JA, Krasnow MA, Metzger RJ. Capillary cell-type specialization in the alveolus. Nature 586: 785–789, 2020. doi: 10.1038/s41586-020-2822-7. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 6. Kalluri AS, Vellarikkal SK, Edelman ER, Nguyen L, Subramanian A, Ellinor PT, Regev A, Kathiresan S, Gupta RM. Single-Cell Analysis of the Normal Mouse Aorta Reveals Functionally Distinct Endothelial Cell Populations. Circulation 140: 147–163, 2019. doi: 10.1161/CIRCULATIONAHA.118.038362. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 7. de Queiroz DB, Parente JM, Pernomian L, Waigi EW, Alfaidi M, Tan W, McCarthy CG, Wenceslau CF. Endothelial Cell Phenotypic Plasticity in Cardiovascular Physiology and Disease: Mechanisms and Therapeutic Prospects. Am J Hypertens 38: 411–421, 2025. doi: 10.1093/ajh/hpaf027. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 8. Ottersbach K. Endothelial-to-haematopoietic transition: an update on the process of making blood. Biochem Soc Trans 47: 591–601, 2019. doi: 10.1042/BST20180320. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 9. Andueza A, Kumar S, Kim J, Kang D-W, Mumme HL, Perez JI, Villa-Roel N, Jo H. Endothelial Reprogramming by Disturbed Flow Revealed by Single-Cell RNA and Chromatin Accessibility Study. Cell Rep 33: 108491, 2020. doi: 10.1016/j.celrep.2020.108491. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 10. Cianciolo G, La Manna G, Della Bella E, Cappuccilli ML, Angelini ML, Dormi A, Capelli I, Laterza C, Costa R, Alviano F, Donati G, Ronco C, Stefoni S. Effect of vitamin D receptor activator therapy on vitamin D receptor and osteocalcin expression in circulating endothelial progenitor cells of hemodialysis patients. Blood Purif 35: 187–95, 2013. doi: 10.1159/000347102. [ DOI ] [ PubMed ] [ Google Scholar ] 11. Zhang H, Wang L, Si D, Wang C, Yang J, Jiang P, Du C, Wang J. Correlation between osteocalcin-positive endothelial progenitor cells and spotty calcification in patients with coronary artery disease. Clin Exp Pharmacol Physiol 42: 734–9, 2015. doi: 10.1111/1440-1681.12366. [ DOI ] [ PubMed ] [ Google Scholar ] 12. Jiang W, Zhang Z, Li Y, Chen C, Yang H, Lin Q, Hu M, Qin X. The Cell Origin and Role of Osteoclastogenesis and Osteoblastogenesis in Vascular Calcification. Front Cardiovasc Med 8: 639740, 2021. doi: 10.3389/fcvm.2021.639740. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 13. Arciniegas E, Frid MG, Douglas IS, Stenmark KR. Perspectives on endothelial-to-mesenchymal transition: potential contribution to vascular remodeling in chronic pulmonary hypertension. Am J Physiol Lung Cell Mol Physiol 293: L1–8, 2007. doi: 10.1152/ajplung.00378.2006. [ DOI ] [ PubMed ] [ Google Scholar ] 14. Pernomian L, Waigi EW, Nguyen V, Mohammed AD, da Costa TJ, Fontes MT, Kubinak JL, Aitken A, Biancardi VC, Sinclair DA, McCarthy CG, Wang Y, Tan W, Wenceslau CF. A Single-Short Partial Reprogramming of the Endothelial Cells decreases Blood Pressure via attenuation of EndMT in Hypertensive Mice. . [ Google Scholar ] 15. Weil BR, Stauffer BL, Greiner JJ, DeSouza CA. Prehypertension is associated with impaired nitric oxide-mediated endothelium-dependent vasodilation in sedentary adults. Am J Hypertens 24: 976–81, 2011. doi: 10.1038/ajh.2011.88. [ DOI ] [ PubMed ] [ Google Scholar ] 16. Rossi R, Chiurlia E, Nuzzo A, Cioni E, Origliani G, Modena MG. Flow-mediated vasodilation and the risk of developing hypertension in healthy postmenopausal women. J Am Coll Cardiol 44: 1636–1640, 2004. doi: 10.1016/j.jacc.2004.07.027. [ DOI ] [ PubMed ] [ Google Scholar ] 17. Taddei S, Virdis A, Ghiadoni L, Salvetti G, Salvetti A. Endothelial dysfunction in hypertension. J Nephrol 13: 205–10, 2000. [ PubMed ] [ Google Scholar ] 18. Bharani A, Jain N, Jain A, Deedwania P. Endothelium-dependent vasodilation is impaired in healthy offspring of hypertensive parents. Indian Heart J 63: 255–8, 2011. [ PubMed ] [ Google Scholar ] 19. von Offenberg Sweeney N, Cummins PM, Birney YA, Cullen JP, Redmond EM, Cahill PA. Cyclic strain-mediated regulation of endothelial matrix metalloproteinase-2 expression and activity. Cardiovasc Res 63: 625–34, 2004. doi: 10.1016/j.cardiores.2004.05.008. [ DOI ] [ PubMed ] [ Google Scholar ] 20. Zhao S, Suciu A, Ziegler T, Moore JE, Bürki E, Meister JJ, Brunner HR. Synergistic effects of fluid shear stress and cyclic circumferential stretch on vascular endothelial cell morphology and cytoskeleton. Arterioscler Thromb Vasc Biol 15: 1781–6, 1995. doi: 10.1161/01.atv.15.10.1781. [ DOI ] [ PubMed ] [ Google Scholar ] 21. Intengan HD, Schiffrin EL. Vascular remodeling in hypertension: roles of apoptosis, inflammation, and fibrosis. Hypertension 38: 581–7, 2001. doi: 10.1161/hy09t1.096249. [ DOI ] [ PubMed ] [ Google Scholar ] 22. Edwards JM, Roy S, Galla SL, Tomcho JC, Bearss NR, Waigi EW, Mell B, Cheng X, Saha P, Vijay-Kumar M, McCarthy CG, Joe B, Wenceslau CF. FPR-1 (Formyl Peptide Receptor-1) Activation Promotes Spontaneous, Premature Hypertension in Dahl Salt-Sensitive Rats. Hypertension 77: 1191–1202, 2021. doi: 10.1161/HYPERTENSIONAHA.120.16237. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 23. Eguchi S, Torimoto K, Adebiyi A, Kanthakumar P, Bomfim GF, Wenceslau CF, Dahlen SA, Osei-Owusu P. Milestone Papers on Signal Transduction Mechanisms of Hypertension and Its Complications. Hypertension 81: 977–990, 2024. doi: 10.1161/HYPERTENSIONAHA.123.21365. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 24. Lim XR, Harraz OF. Mechanosensing by Vascular Endothelium. Annu Rev Physiol 86: 71–97, 2024. doi: 10.1146/annurev-physiol-042022-030946. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 25. Garland CJ, Plane F, Kemp BK, Cocks TM. Endothelium-dependent hyperpolarization: a role in the control of vascular tone. Trends Pharmacol Sci 16: 23–30, 1995. doi: 10.1016/s0165-6147(00)88969-5. [ DOI ] [ PubMed ] [ Google Scholar ] 26. Dhaun N, Goddard J, Kohan DE, Pollock DM, Schiffrin EL, Webb DJ. Role of endothelin-1 in clinical hypertension: 20 years on. Hypertension 52: 452–9, 2008. doi: 10.1161/HYPERTENSIONAHA.108.117366. [ DOI ] [ PubMed ] [ Google Scholar ] 27. Wong MS- K, Vanhoutte PM. COX-mediated endothelium-dependent contractions: from the past to recent discoveries. Acta Pharmacol Sin 31: 1095–102, 2010. doi: 10.1038/aps.2010.127. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 28. Xavier FE, Aras-López R, Arroyo-Villa I, Del Campo L, Salaices M, Rossoni LV, Ferrer M, Balfagón G. Aldosterone induces endothelial dysfunction in resistance arteries from normotensive and hypertensive rats by increasing thromboxane A 2 and prostacyclin. Br J Pharmacol 154: 1225–1235, 2008. doi: 10.1038/bjp.2008.200. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 29. Avvisato R, Jankauskas SS, Varzideh F, Kansakar U, Mone P, Santulli G. Sortilin and hypertension. Curr Opin Nephrol Hypertens 32: 134–140, 2023. doi: 10.1097/MNH.0000000000000866. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 30. Di Pietro P, Carrizzo A, Sommella E, Oliveti M, Iacoviello L, Di Castelnuovo A, Acernese F, Damato A, De Lucia M, Merciai F, Iesu P, Venturini E, Izzo R, Trimarco V, Ciccarelli M, Giugliano G, Carnevale R, Cammisotto V, Migliarino S, Virtuoso N, Strianese A, Izzo V, Campiglia P, Ciaglia E, Levkau B, Puca AA, Vecchione C. Targeting the ASMase/S1P pathway protects from sortilin-evoked vascular damage in hypertension. J Clin Invest 132, 2022. doi: 10.1172/JCI146343. [ DOI ] [ Google Scholar ] 31. Freed JK, Beyer AM, LoGiudice JA, Hockenberry JC, Gutterman DD. Ceramide changes the mediator of flow-induced vasodilation from nitric oxide to hydrogen peroxide in the human microcirculation. Circ Res 115: 525–32, 2014. doi: 10.1161/CIRCRESAHA.115.303881. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 32. Vischer UM, Jornot L, Wollheim CB, Theler JM. Reactive oxygen intermediates induce regulated secretion of von Willebrand factor from cultured human vascular endothelial cells. Blood 85: 3164–72, 1995. [ PubMed ] [ Google Scholar ] 33. Lip GY, Blann A. von Willebrand factor: a marker of endothelial dysfunction in vascular disorders? Cardiovasc Res 34: 255–65, 1997. doi: 10.1016/s0008-6363(97)00039-4. [ DOI ] [ PubMed ] [ Google Scholar ] 34. Martin FA, Murphy RP, Cummins PM. Thrombomodulin and the vascular endothelium: insights into functional, regulatory, and therapeutic aspects. Am J Physiol Heart Circ Physiol 304: H1585–97, 2013. doi: 10.1152/ajpheart.00096.2013. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 35. Hrafnkelsdóttir T, Ottosson P, Gudnason T, Samuelsson O, Jern S. Impaired endothelial release of tissue-type plasminogen activator in patients with chronic kidney disease and hypertension. Hypertension 44: 300–4, 2004. doi: 10.1161/01.HYP.0000137380.91476.fb. [ DOI ] [ PubMed ] [ Google Scholar ] 36. Dejana E, Orsenigo F, Lampugnani MG. The role of adherens junctions and VE-cadherin in the control of vascular permeability. J Cell Sci 121: 2115–22, 2008. doi: 10.1242/jcs.017897. [ DOI ] [ PubMed ] [ Google Scholar ] 37. Zhang X, Sun D, Song JW, Zullo J, Lipphardt M, Coneh-Gould L, Goligorsky MS. Endothelial cell dysfunction and glycocalyx - A vicious circle. Matrix Biol 71–72: 421–431, 2018. doi: 10.1016/j.matbio.2018.01.026. [ DOI ] [ Google Scholar ] 38. McCarthy CG, Goulopoulou S, Wenceslau CF, Spitler K, Matsumoto T, Webb RC. Toll-like receptors and damage-associated molecular patterns: novel links between inflammation and hypertension. Am J Physiol Heart Circ Physiol 306: H184–96, 2014. doi: 10.1152/ajpheart.00328.2013. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 39. McMullan RR, McAuley DF, O’Kane CM, Silversides JA. Vascular leak in sepsis: physiological basis and potential therapeutic advances. Crit Care 28: 97, 2024. doi: 10.1186/s13054-024-04875-6. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 40. De Sanctis F, Ugel S, Facciponte J, Facciabene A. The dark side of tumor-associated endothelial cells. Semin Immunol 35: 35–47, 2018. doi: 10.1016/j.smim.2018.02.002. [ DOI ] [ PubMed ] [ Google Scholar ] 41. Ebeling S, Kowalczyk A, Perez-Vazquez D, Mattiola I. Regulation of tumor angiogenesis by the crosstalk between innate immunity and endothelial cells. Front Oncol 13: 1171794, 2023. doi: 10.3389/fonc.2023.1171794. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 42. Ziche M, Morbidelli L, Choudhuri R, Zhang HT, Donnini S, Granger HJ, Bicknell R. Nitric oxide synthase lies downstream from vascular endothelial growth factor-induced but not basic fibroblast growth factor-induced angiogenesis. J Clin Invest 99: 2625–34, 1997. doi: 10.1172/JCI119451. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 43. Antonetti DA, Klein R, Gardner TW. Diabetic retinopathy. N Engl J Med 366: 1227–39, 2012. doi: 10.1056/NEJMra1005073. [ DOI ] [ PubMed ] [ Google Scholar ] 44. Yamagishi S, Maeda S, Matsui T, Ueda S, Fukami K, Okuda S. Role of advanced glycation end products (AGEs) and oxidative stress in vascular complications in diabetes. Biochim Biophys Acta 1820: 663–71, 2012. doi: 10.1016/j.bbagen.2011.03.014. [ DOI ] [ PubMed ] [ Google Scholar ] 45. Pacher P, Szabó C. Role of peroxynitrite in the pathogenesis of cardiovascular complications of diabetes. Curr Opin Pharmacol 6: 136–41, 2006. doi: 10.1016/j.coph.2006.01.001. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 46. Ceriello A, Mercuri F, Quagliaro L, Assaloni R, Motz E, Tonutti L, Taboga C. Detection of nitrotyrosine in the diabetic plasma: evidence of oxidative stress. Diabetologia 44: 834–8, 2001. doi: 10.1007/s001250100529. [ DOI ] [ PubMed ] [ Google Scholar ] 47. Carreau A, El Hafny-Rahbi B, Matejuk A, Grillon C, Kieda C. Why is the partial oxygen pressure of human tissues a crucial parameter? Small molecules and hypoxia. J Cell Mol Med 15: 1239–53, 2011. doi: 10.1111/j.1582-4934.2011.01258.x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 48. Attaye I, Smulders YM, de Waard MC, Oudemans-van Straaten HM, Smit B, Van Wijhe MH, Musters RJ, Koolwijk P, Spoelstra-de Man AME. The effects of hyperoxia on microvascular endothelial cell proliferation and production of vaso-active substances. Intensive Care Med Exp 5: 22, 2017. doi: 10.1186/s40635-017-0135-4. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 49. Uno K, Merges CA, Grebe R, Lutty GA, Prow TW. Hyperoxia inhibits several critical aspects of vascular development. Dev Dyn 236: 981–90, 2007. doi: 10.1002/dvdy.21122. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 50. Rubanyi GM, Vanhoutte PM. Superoxide anions and hyperoxia inactivate endothelium-derived relaxing factor. Am J Physiol 250: H822–7, 1986. doi: 10.1152/ajpheart.1986.250.5.H822. [ DOI ] [ PubMed ] [ Google Scholar ] 51. Wrobeln A, Schlüter KD, Linders J, Zähres M, Mayer C, Kirsch M, Ferenz KB. Functionality of albumin-derived perfluorocarbon-based artificial oxygen carriers in the Langendorff-heart †. Artif Cells Nanomed Biotechnol 45: 723–730, 2017. doi: 10.1080/21691401.2017.1284858. [ DOI ] [ PubMed ] [ Google Scholar ] 52. Rasmussen JK, Boedtkjer E. Carbonic anhydrase inhibitors modify intracellular pH transients and contractions of rat middle cerebral arteries during CO2/HCO3- fluctuations. J Cereb Blood Flow Metab 38: 492–505, 2018. doi: 10.1177/0271678X17699224. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 53. Hansen KB, Staehr C, Rohde PD, Homilius C, Kim S, Nyegaard M, Matchkov VV, Boedtkjer E. PTPRG is an ischemia risk locus essential for HCO3--dependent regulation of endothelial function and tissue perfusion. Elife 9, 2020. doi: 10.7554/eLife.57553. [ DOI ] [ Google Scholar ] 54. Schmitz B, Nedele J, Guske K, Maase M, Lenders M, Schelleckes M, Kusche-Vihrog K, Brand S-M, Brand E. Soluble adenylyl cyclase in vascular endothelium: gene expression control of epithelial sodium channel-α, Na+/K+-ATPase-α/β, and mineralocorticoid receptor. Hypertension 63: 753–61, 2014. doi: 10.1161/HYPERTENSIONAHA.113.02061. [ DOI ] [ PubMed ] [ Google Scholar ] 55. Boedtkjer E, Hansen KB, Boedtkjer DMB, Aalkjaer C, Boron WF. Extracellular HCO3- is sensed by mouse cerebral arteries: Regulation of tone by receptor protein tyrosine phosphatase γ. J Cereb Blood Flow Metab 36: 965–80, 2016. doi: 10.1177/0271678X15610787. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 56. Goodrich HR, Berry AA, Montgomery DW, Davison WG, Wilson RW. Fish feeds supplemented with calcium-based buffering minerals decrease stomach acidity, increase the blood alkaline tide and cost more to digest. Sci Rep 12: 18468, 2022. doi: 10.1038/s41598-022-22496-3. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 57. Stabenau EKHT. Determination of the constants of the Henderson–Hasselbalch Equation, αCO2 and pKa, in sea turtle plasma. . J Exp Bio 180: 311–314, 1993. [ Google Scholar ] 58. Boedtkjer E, Aalkjaer C. The solution to bicarbonate. Am J Physiol Heart Circ Physiol 322: H685–H686, 2022. doi: 10.1152/ajpheart.00057.2022. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 59. Turlapaty PD, Altura BT, Altura BM. Interactions of Tris buffer and ethanol on agonist-induced responses of vascular smooth muscle and on calcium-45 uptake. J Pharmacol Exp Ther 211: 59–67, 1979. [ PubMed ] [ Google Scholar ] 60. Juel C. Changes in interstitial K+ and pH during exercise: implications for blood flow regulation. Appl Physiol Nutr Metab 32: 846–51, 2007. doi: 10.1139/H07-065. [ DOI ] [ PubMed ] [ Google Scholar ] 61. Warner MR, Kroeker TS, Zipes DP. Sympathetic stimulation and norepinephrine infusion modulate extracellular potassium concentration during acute myocardial ischemia. Circ Res 71: 1078–87, 1992. doi: 10.1161/01.res.71.5.1078. [ DOI ] [ PubMed ] [ Google Scholar ] 62. McKercher HG, Derewlany LO, Radde IC. Free calcium concentrations in Krebs-Ringer bicarbonate buffer: effects on 45Ca- and 32P-transport across the perfused guinea pig placenta. Biochem Biophys Res Commun 105: 841–6, 1982. doi: 10.1016/0006-291x(82)91046-4. [ DOI ] [ PubMed ] [ Google Scholar ] 63. Clyne AM. Endothelial response to glucose: dysfunction, metabolism, and transport. Biochem Soc Trans 49: 313–325, 2021. doi: 10.1042/BST20200611. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 64. Ashbrook JD, Spector AA, Santos EC, Fletcher JE. Long chain fatty acid binding to human plasma albumin. J Biol Chem 250: 2333–8, 1975. [ PubMed ] [ Google Scholar ] 65. MacKenzie A, Wadsworth RM. Extracellular L-arginine is required for optimal NO synthesis by eNOS and iNOS in the rat mesenteric artery wall. Br J Pharmacol 139: 1487–97, 2003. doi: 10.1038/sj.bjp.0705380. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 66. Park S-Y, Ives SJ, Gifford JR, Andtbacka RHI, Hyngstrom JR, Reese V, Layec G, Bharath LP, Symons JD, Richardson RS. Impact of age on the vasodilatory function of human skeletal muscle feed arteries. Am J Physiol Heart Circ Physiol 310: H217–25, 2016. doi: 10.1152/ajpheart.00716.2015. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 67. Nedergaard OA, Vagne A, Bevan JA. Effect of the chelating agents, EDTA, 2,2’-bipyridine, 8-hydroxyquinoline and pyrophosphoric acid, on norepinephrine uptake by rabbit aorta. J Pharmacol Exp Ther 163: 136–46, 1968. [ PubMed ] [ Google Scholar ] 68. Thbayh DK, Palusiak M, Viskolcz B, Fiser B. Comparative study of the antioxidant capability of EDTA and Irganox. Heliyon 9: e16064, 2023. doi: 10.1016/j.heliyon.2023.e16064. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 69. Bukoski RD, Shearin S, Jackson WF, Pamarthi MF. Inhibition of Ca2+-induced relaxation by oxidized tungsten wires and paratungstate. J Pharmacol Exp Ther 299: 343–50, 2001. [ PubMed ] [ Google Scholar ] 70. Vanhoutte PM, Zhao Y, Xu A, Leung SWS. Thirty Years of Saying NO: Sources, Fate, Actions, and Misfortunes of the Endothelium-Derived Vasodilator Mediator. Circ Res 119: 375–96, 2016. doi: 10.1161/CIRCRESAHA.116.306531. [ DOI ] [ PubMed ] [ Google Scholar ] 71. Chataigneau T, Félétou M, Huang PL, Fishman MC, Duhault J, Vanhoutte PM. Acetylcholine-induced relaxation in blood vessels from endothelial nitric oxide synthase knockout mice. Br J Pharmacol 126: 219–26, 1999. doi: 10.1038/sj.bjp.0702300. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 72. Shimokawa H, Yasutake H, Fujii K, Owada MK, Nakaike R, Fukumoto Y, Takayanagi T, Nagao T, Egashira K, Fujishima M, Takeshita A. The importance of the hyperpolarizing mechanism increases as the vessel size decreases in endothelium-dependent relaxations in rat mesenteric circulation. J Cardiovasc Pharmacol 28: 703–11, 1996. doi: 10.1097/00005344-199611000-00014. [ DOI ] [ PubMed ] [ Google Scholar ] 73. Lüscher TF, Vanhoutte PM. Endothelium-dependent contractions to acetylcholine in the aorta of the spontaneously hypertensive rat. Hypertension 8: 344–8, 1986. doi: 10.1161/01.hyp.8.4.344. [ DOI ] [ PubMed ] [ Google Scholar ] 74. Roque FR, Briones AM, García-Redondo AB, Galán M, Martínez-Revelles S, Avendaño MS, Cachofeiro V, Fernandes T, Vassallo DV, Oliveira EM, Salaices M. Aerobic exercise reduces oxidative stress and improves vascular changes of small mesenteric and coronary arteries in hypertension. Br J Pharmacol 168: 686–703, 2013. doi: 10.1111/j.1476-5381.2012.02224.x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 75. Iwamoto T. Vascular Na+/Ca2+ exchanger: implications for the pathogenesis and therapy of salt-dependent hypertension. Am J Physiol Regul Integr Comp Physiol 290: R536–45, 2006. doi: 10.1152/ajpregu.00592.2005. [ DOI ] [ PubMed ] [ Google Scholar ] 76. Wanstall JC, Jeffery TK, Gambino A, Lovren F, Triggle CR. Vascular smooth muscle relaxation mediated by nitric oxide donors: a comparison with acetylcholine, nitric oxide and nitroxyl ion. Br J Pharmacol 134: 463–72, 2001. doi: 10.1038/sj.bjp.0704269. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 77. Guizoni DM, Dorighello GG, Oliveira HCF, Delbin MA, Krieger MH, Davel AP. Aerobic exercise training protects against endothelial dysfunction by increasing nitric oxide and hydrogen peroxide production in LDL receptor-deficient mice. J Transl Med 14: 213, 2016. doi: 10.1186/s12967-016-0972-z. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 78. Taddei S, Vanhoutte PM. Role of endothelium in endothelin-evoked contractions in the rat aorta. Hypertension 21: 9–15, 1993. doi: 10.1161/01.hyp.21.1.9. [ DOI ] [ PubMed ] [ Google Scholar ] 79. Zhou Y, Varadharaj S, Zhao X, Parinandi N, Flavahan NA, Zweier JL. Acetylcholine causes endothelium-dependent contraction of mouse arteries. Am J Physiol Heart Circ Physiol 289: H1027–32, 2005. doi: 10.1152/ajpheart.00226.2005. [ DOI ] [ PubMed ] [ Google Scholar ] 80. Privistirescu AI, Sima A, Duicu OM, Timar R, Roșca MG, Sturza A, Muntean DM. Methylene blue alleviates endothelial dysfunction and reduces oxidative stress in aortas from diabetic rats. Can J Physiol Pharmacol 96: 1012–1016, 2018. doi: 10.1139/cjpp-2018-0119. [ DOI ] [ PubMed ] [ Google Scholar ] 81. Dănilă MD, Privistirescu A, Duicu OM, Rațiu CD, Angoulvant D, Muntean DM, Sturza A. The effect of purinergic signaling via the P2Y11 receptor on vascular function in a rat model of acute inflammation. Mol Cell Biochem 431: 37–44, 2017. doi: 10.1007/s11010-017-2973-5. [ DOI ] [ PubMed ] [ Google Scholar ] 82. Mian KB, Martin W. Differential sensitivity of basal and acetylcholine-stimulated activity of nitric oxide to destruction by superoxide anion in rat aorta. Br J Pharmacol 115: 993–1000, 1995. doi: 10.1111/j.1476-5381.1995.tb15909.x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 83. Davel APC, Kawamoto EM, Scavone C, Vassallo DV, Rossoni LV. Changes in vascular reactivity following administration of isoproterenol for 1 week: a role for endothelial modulation. Br J Pharmacol 148: 629–39, 2006. doi: 10.1038/sj.bjp.0706749. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 84. Overend J, Martin W. Differential effects of nitric oxide synthase inhibitors on endothelium-dependent and nitrergic nerve-mediated vasodilatation in the bovine ciliary artery. Br J Pharmacol 150: 488–93, 2007. doi: 10.1038/sj.bjp.0707113. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 85. Archer SL, Hampl V. NG-monomethyl-L-arginine causes nitric oxide synthesis in isolated arterial rings: trouble in paradise. Biochem Biophys Res Commun 188: 590–6, 1992. doi: 10.1016/0006-291x(92)91097-a. [ DOI ] [ PubMed ] [ Google Scholar ] 86. Frew JD, Paisley K, Martin W. Selective inhibition of basal but not agonist-stimulated activity of nitric oxide in rat aorta by NG-monomethyl-L-arginine. Br J Pharmacol 110: 1003–8, 1993. doi: 10.1111/j.1476-5381.1993.tb13913.x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 87. Plane F, Wiley KE, Jeremy JY, Cohen RA, Garland CJ. Evidence that different mechanisms underlie smooth muscle relaxation to nitric oxide and nitric oxide donors in the rabbit isolated carotid artery. Br J Pharmacol 123: 1351–8, 1998. doi: 10.1038/sj.bjp.0701746. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 88. Virdis A, Colucci R, Fornai M, Duranti E, Giannarelli C, Bernardini N, Segnani C, Ippolito C, Antonioli L, Blandizzi C, Taddei S, Salvetti A, Del Tacca M. Cyclooxygenase-1 is involved in endothelial dysfunction of mesenteric small arteries from angiotensin II-infused mice. Hypertension 49: 679–86, 2007. doi: 10.1161/01.HYP.0000253085.56217.11. [ DOI ] [ PubMed ] [ Google Scholar ] 89. Leuranguer V, Gluais P, Vanhoutte PM, Verbeuren TJ, Félétou M. Openers of calcium-activated potassium channels and endothelium-dependent hyperpolarizations in the guinea pig carotid artery. Naunyn Schmiedebergs Arch Pharmacol 377: 101–9, 2008. doi: 10.1007/s00210-008-0267-x. [ DOI ] [ PubMed ] [ Google Scholar ] 90. Juguilon C, Wang Z, Wang Y, Enrick M, Jamaiyar A, Xu Y, Gadd J, Chen C- LW, Pu A, Kolz C, Ohanyan V, Chen Y-R, Hardwick J, Zhang Y, Chilian WM, Yin L. Mechanism of the switch from NO to H2O2 in endothelium-dependent vasodilation in diabetes. Basic Res Cardiol 117: 2, 2022. doi: 10.1007/s00395-022-00910-1. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 91. Celermajer DS, Sorensen KE, Gooch VM, Spiegelhalter DJ, Miller OI, Sullivan ID, Lloyd JK, Deanfield JE. Non-invasive detection of endothelial dysfunction in children and adults at risk of atherosclerosis. Lancet 340: 1111–5, 1992. doi: 10.1016/0140-6736(92)93147-f. [ DOI ] [ PubMed ] [ Google Scholar ] 92. Clifford PS, Madden JA, Hamann JJ, Buckwalter JB, Valic Z. Absence of flow-mediated vasodilation in the rabbit femoral artery. Physiol Res 59: 331–338, 2010. doi: 10.33549/physiolres.931672. [ DOI ] [ PubMed ] [ Google Scholar ] 93. Tesfamariam B, Halpern W. Modulation of adrenergic responses in pressurized resistance arteries by flow. Am J Physiol 253: H1112–9, 1987. doi: 10.1152/ajpheart.1987.253.5.H1112. [ DOI ] [ PubMed ] [ Google Scholar ] 94. Kuo L, Davis MJ, Chilian WM. Endothelium-dependent, flow-induced dilation of isolated coronary arterioles. Am J Physiol 259: H1063–70, 1990. doi: 10.1152/ajpheart.1990.259.4.H1063. [ DOI ] [ PubMed ] [ Google Scholar ] 95. SenthilKumar G, Katunaric B, Bordas-Murphy H, Young M, Doren EL, Schulz ME, Widlansky ME, Freed JK. 17β-Estradiol promotes sex-specific dysfunction in isolated human arterioles. Am J Physiol Heart Circ Physiol 324: H330–H337, 2023. doi: 10.1152/ajpheart.00708.2022. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 96. Hader SN, Zinkevich N, Norwood Toro LE, Kriegel AJ, Kong A, Freed JK, Gutterman DD, Beyer AM. Detrimental effects of chemotherapy on human coronary microvascular function. Am J Physiol Heart Circ Physiol 317: H705–H710, 2019. doi: 10.1152/ajpheart.00370.2019. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 97. Durand MJ, Dharmashankar K, Bian J-T, Das E, Vidovich M, Gutterman DD, Phillips SA. Acute exertion elicits a H2O2-dependent vasodilator mechanism in the microvasculature of exercise-trained but not sedentary adults. Hypertension 65: 140–5, 2015. doi: 10.1161/HYPERTENSIONAHA.114.04540. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 98. Shimoda LA, Norins NA, Jeutter DC, Madden JA. Flow-induced responses in piglet isolated cerebral arteries. Pediatr Res 39: 574–83, 1996. doi: 10.1203/00006450-199604000-00002. [ DOI ] [ PubMed ] [ Google Scholar ] 99. Tribe RM, Thomas CR, Poston L. Flow-induced dilatation in isolated resistance arteries from control and streptozotocin-diabetic rats. Diabetologia 41: 34–9, 1998. doi: 10.1007/s001250050863. [ DOI ] [ PubMed ] [ Google Scholar ] 100. Garcia-Roldan JL, Bevan JA. Flow-induced constriction and dilation of cerebral resistance arteries. Circ Res 66: 1445–8, 1990. doi: 10.1161/01.res.66.5.1445. [ DOI ] [ PubMed ] [ Google Scholar ] 101. Cohen KE, Katunaric B, Schulz ME, SenthilKumar G, Young MS, Mace JE, Freed JK. Role of Adiponectin Receptor 1 in Promoting Nitric Oxide-Mediated Flow-Induced Dilation in the Human Microvasculature. Front Pharmacol 13: 875900, 2022. doi: 10.3389/fphar.2022.875900. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 102. Kadlec AO, Chabowski DS, Ait-Aissa K, Hockenberry JC, Otterson MF, Durand MJ, Freed JK, Beyer AM, Gutterman DD. PGC-1α (Peroxisome Proliferator-Activated Receptor γ Coactivator 1-α) Overexpression in Coronary Artery Disease Recruits NO and Hydrogen Peroxide During Flow-Mediated Dilation and Protects Against Increased Intraluminal Pressure. Hypertension 70: 166–173, 2017. doi: 10.1161/HYPERTENSIONAHA.117.09289. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 103. Wolpe AG, Ruddiman CA, Hall PJ, Isakson BE. Polarized Proteins in Endothelium and Their Contribution to Function. J Vasc Res 58: 65–91, 2021. doi: 10.1159/000512618. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 104. Sonkusare SK, Bonev AD, Ledoux J, Liedtke W, Kotlikoff MI, Heppner TJ, Hill-Eubanks DC, Nelson MT. Elementary Ca2+ signals through endothelial TRPV4 channels regulate vascular function. Science 336: 597–601, 2012. doi: 10.1126/science.1216283. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 105. Luse MA, Schug WJ, Dunaway LS, Nyshadham S, Loeb SA, Carvalho A, Tessema R, Pavelec C, Keller TCS, Shu X, Ruddiman CA, Kosmach A, Sveeggen TM, Mitchell R, Bagher P, Minshall RD, Leitnger N, Columbus L, Levental KR, Levental I, Cortese-Krott M, Isakson BE. Nitrosation of CD36 Regulates Endothelial Function and Serum Lipids. Arterioscler Thromb Vasc Biol 45: 1067–1086, 2025. doi: 10.1161/ATVBAHA.124.321964. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 106. Maier-Begandt D, Comstra HS, Molina SA, Krüger N, Ruddiman CA, Chen Y-L, Chen X, Biwer LA, Johnstone SR, Lohman AW, Good ME, DeLalio LJ, Hong K, Bacon HM, Yan Z, Sonkusare SK, Koval M, Isakson BE. A venous-specific purinergic signaling cascade initiated by Pannexin 1 regulates TNFα-induced increases in endothelial permeability. Sci Signal 14, 2021. doi: 10.1126/scisignal.aba2940. [ DOI ] [ Google Scholar ] 107. Boedtkjer E, Aalkjaer C. Insulin inhibits Na+/H+ exchange in vascular smooth muscle and endothelial cells in situ: involvement of H2O2 and tyrosine phosphatase SHP-2. Am J Physiol Heart Circ Physiol 296: H247–55, 2009. doi: 10.1152/ajpheart.00725.2008. [ DOI ] [ PubMed ] [ Google Scholar ] 108. Krogh A, Harrop GA, Rehberg PB. Studies on the physiology of capillaries: III. The innervation of the blood vessels in the hind legs of the frog. J Physiol 56: 179–89, 1922. doi: 10.1113/jphysiol.1922.sp002000. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 109. Duling BR, Berne RM. Propagated vasodilation in the microcirculation of the hamster cheek pouch. Circ Res 26: 163–70, 1970. doi: 10.1161/01.res.26.2.163. [ DOI ] [ PubMed ] [ Google Scholar ] 110. Segal SS, Duling BR. Flow control among microvessels coordinated by intercellular conduction. Science 234: 868–70, 1986. doi: 10.1126/science.3775368. [ DOI ] [ PubMed ] [ Google Scholar ] 111. Rhodin JA. The ultrastructure of mammalian arterioles and precapillary sphincters. J Ultrastruct Res 18: 181–223, 1967. doi: 10.1016/s0022-5320(67)80239-9. [ DOI ] [ PubMed ] [ Google Scholar ] 112. Dora KA, Doyle MP, Duling BR. Elevation of intracellular calcium in smooth muscle causes endothelial cell generation of NO in arterioles. Proc Natl Acad Sci U S A 94: 6529–34, 1997. doi: 10.1073/pnas.94.12.6529. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 113. Sandow SL, Hill CE. Incidence of myoendothelial gap junctions in the proximal and distal mesenteric arteries of the rat is suggestive of a role in endothelium-derived hyperpolarizing factor-mediated responses. Circ Res 86: 341–6, 2000. doi: 10.1161/01.res.86.3.341. [ DOI ] [ PubMed ] [ Google Scholar ] 114. Yashiro Y, Duling BR. Integrated Ca(2+) signaling between smooth muscle and endothelium of resistance vessels. Circ Res 87: 1048–54, 2000. doi: 10.1161/01.res.87.11.1048. [ DOI ] [ PubMed ] [ Google Scholar ] 115. Emerson GG, Segal SS. Endothelial cell pathway for conduction of hyperpolarization and vasodilation along hamster feed artery. Circ Res 86: 94–100, [date unknown]. doi: 10.1161/01.res.86.1.94. [ DOI ] [ Google Scholar ] 116. Emerson GG, Segal SS. Electrical coupling between endothelial cells and smooth muscle cells in hamster feed arteries: role in vasomotor control. Circ Res 87: 474–9, 2000. doi: 10.1161/01.res.87.6.474. [ DOI ] [ PubMed ] [ Google Scholar ] 117. Segal SS, Jacobs TL. Role for endothelial cell conduction in ascending vasodilatation and exercise hyperaemia in hamster skeletal muscle. J Physiol 536: 937–46, 2001. doi: 10.1111/j.1469-7793.2001.00937.x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 118. Tallini YN, Brekke JF, Shui B, Doran R, Hwang S, Nakai J, Salama G, Segal SS, Kotlikoff MI. Propagated endothelial Ca2+ waves and arteriolar dilation in vivo: measurements in Cx40BAC GCaMP2 transgenic mice. Circ Res 101: 1300–9, 2007. doi: 10.1161/CIRCRESAHA.107.149484. [ DOI ] [ PubMed ] [ Google Scholar ] 119. Domeier TL, Segal SS. Electromechanical and pharmacomechanical signalling pathways for conducted vasodilatation along endothelium of hamster feed arteries. J Physiol 579: 175–86, 2007. doi: 10.1113/jphysiol.2006.124529. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 120. Garland CJ, Bagher P, Powell C, Ye X, Lemmey HAL, Borysova L, Dora KA. Voltage-dependent Ca2+ entry into smooth muscle during contraction promotes endothelium-mediated feedback vasodilation in arterioles. Sci Signal 10, 2017. doi: 10.1126/scisignal.aal3806. [ DOI ] [ Google Scholar ] 121. Isakson BE, Duling BR. Heterocellular contact at the myoendothelial junction influences gap junction organization. Circ Res 97: 44–51, 2005. doi: 10.1161/01.RES.0000173461.36221.2e. [ DOI ] [ PubMed ] [ Google Scholar ] 122. Ledoux J, Taylor MS, Bonev AD, Hannah RM, Solodushko V, Shui B, Tallini Y, Kotlikoff MI, Nelson MT. Functional architecture of inositol 1,4,5-trisphosphate signaling in restricted spaces of myoendothelial projections. Proc Natl Acad Sci U S A 105: 9627–32, 2008. doi: 10.1073/pnas.0801963105. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 123. Tran CHT, Taylor MS, Plane F, Nagaraja S, Tsoukias NM, Solodushko V, Vigmond EJ, Furstenhaupt T, Brigdan M, Welsh DG. Endothelial Ca2+ wavelets and the induction of myoendothelial feedback. Am J Physiol Cell Physiol 302: C1226–42, 2012. doi: 10.1152/ajpcell.00418.2011. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 124. Afshar Y, Ma F, Quach A, Jeong A, Sunshine HL, Freitas V, Jami-Alahmadi Y, Helaers R, Li X, Pellegrini M, Wohlschlegel JA, Romanoski CE, Vikkula M, Iruela-Arispe ML. Transcriptional drifts associated with environmental changes in endothelial cells. Elife 12, 2023. doi: 10.7554/eLife.81370. [ DOI ] [ Google Scholar ] 125. Socha MJ, Hakim CH, Jackson WF, Segal SS. Temperature effects on morphological integrity and Ca 2+ signaling in freshly isolated murine feed artery endothelial cell tubes. Am J Physiol Heart Circ Physiol 301: H773–83, 2011. doi: 10.1152/ajpheart.00214.2011. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 126. Socha MJ, Segal SS. Isolation of microvascular endothelial tubes from mouse resistance arteries. J Vis Exp : e50759, 2013. doi: 10.3791/50759. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 127. Behringer EJ, Segal SS. Tuning electrical conduction along endothelial tubes of resistance arteries through Ca(2+)-activated K(+) channels. Circ Res 110: 1311–21, 2012. doi: 10.1161/CIRCRESAHA.111.262592. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 128. Norton CE, Segal SS. Calcitonin gene-related peptide hyperpolarizes mouse pulmonary artery endothelial tubes through KATP channel activation. Am J Physiol Lung Cell Mol Physiol 315: L212–L226, 2018. doi: 10.1152/ajplung.00044.2018. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 129. Behringer EJ, Shaw RL, Westcott EB, Socha MJ, Segal SS. Aging impairs electrical conduction along endothelium of resistance arteries through enhanced Ca2+-activated K+ channel activation. Arterioscler Thromb Vasc Biol 33: 1892–901, 2013. doi: 10.1161/ATVBAHA.113.301514. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 130. Socha MJ, Boerman EM, Behringer EJ, Shaw RL, Domeier TL, Segal SS. Advanced age protects microvascular endothelium from aberrant Ca(2+) influx and cell death induced by hydrogen peroxide. J Physiol 593: 2155–69, 2015. doi: 10.1113/JP270169. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 131. Hakim CH, Jackson WF, Segal SS. Connexin isoform expression in smooth muscle cells and endothelial cells of hamster cheek pouch arterioles and retractor feed arteries. Microcirculation 15: 503–14, 2008. doi: 10.1080/10739680801982808. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 132. Garcia SM, Naik JS, Resta TC, Jernigan NL. Acid-sensing ion channel 1a activates IKCa/SKCa channels and contributes to endothelium-dependent dilation. J Gen Physiol 155, 2023. doi: 10.1085/jgp.202213173. [ DOI ] [ Google Scholar ] 133. Power G, Lateef OM, Ramirez-Perez FI, Lazo-Fernandez Y, Augenreich MA, Ferreira-Santos L, Soares RN, Gonzalez-Vallejo JD, Morales-Quinones M, Norton CE, Manrique-Acevedo C, Martinez-Lemus LA, Padilla J. Reduced cofilin activity as a mechanism contributing to endothelial cell stiffening in type 2 diabetes. Am J Physiol Heart Circ Physiol 328: H84–H92, 2025. doi: 10.1152/ajpheart.00667.2024. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 134. Hakim MA, Behringer EJ. Development of Alzheimer’s Disease Progressively Alters Sex-Dependent KCa and Sex-Independent KIR Channel Function in Cerebrovascular Endothelium. J Alzheimers Dis 76: 1423–1442, 2020. doi: 10.3233/JAD-200085. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 135. Polk FD, Hakim MA, Silva JF, Behringer EJ, Pires PW. Endothelial KIR2 channel dysfunction in aged cerebral parenchymal arterioles. Am J Physiol Heart Circ Physiol 325: H1360–72, 2023. doi: 10.1152/ajpheart.00279.2023. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 136. Norton CE, Shaw RL, Safa Dockery B, Domeier TL, Segal SS. Advanced age and female sex protect cerebral arteries from mitochondrial depolarization and apoptosis during acute oxidative stress. Aging Cell 23: e14110, 2024. doi: 10.1111/acel.14110. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 137. Behringer EJ, Scallan JP, Jafarnejad M, Castorena-Gonzalez JA, Zawieja SD, Moore JE, Davis MJ, Segal SS. Calcium and electrical dynamics in lymphatic endothelium. J Physiol 595: 7347–7368, 2017. doi: 10.1113/JP274842. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 138. Townsley MI. Structure and composition of pulmonary arteries, capillaries, and veins. Compr Physiol 2: 675–709, 2012. doi: 10.1002/cphy.c100081. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 139. Socha MJ, Domeier TL, Behringer EJ, Segal SS. Coordination of intercellular Ca(2+) signaling in endothelial cell tubes of mouse resistance arteries. Microcirculation 19: 757–70, 2012. doi: 10.1111/micc.12000. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 140. Wilson C, Lee MD, Buckley C, Zhang X, McCarron JG. Mitochondrial ATP Production is Required for Endothelial Cell Control of Vascular Tone. Function (Oxf) 4: zqac063, 2023. doi: 10.1093/function/zqac063. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 141. Kavoussi PK, Heberlein K, Straub AC, Lowe GJ, Oliver JL, Smith RP, Steers WD, Annex BH, Isakson BE, Lysiak JJ. Recombinant PAI-1 therapy restores myoendothelial junctions and erectile function in PAI-1-deficient mice. Andrologia 47: 1147–52, 2015. doi: 10.1111/and.12395. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 142. Ruddiman CA, Peckham R, Luse MA, Chen Y-L, Kuppusamy M, Corliss BA, Hall PJ, Lin C-J, Peirce SM, Sonkusare SK, Mecham RP, Wagenseil JE, Isakson BE. Polarized localization of phosphatidylserine in the endothelium regulates Kir2.1. JCI Insight 8, 2023. doi: 10.1172/jci.insight.165715. [ DOI ] [ Google Scholar ] 143. Sargent SM, Bonney SK, Li Y, Stamenkovic S, Takeno MM, Coelho-Santos V, Shih AY. Endothelial structure contributes to heterogeneity in brain capillary diameter. Vasc Biol 5, 2023. doi: 10.1530/VB-23-0010. [ DOI ] [ Google Scholar ] 144. Dora KA, Sandow SL, Gallagher NT, Takano H, Rummery NM, Hill CE, Garland CJ. Myoendothelial gap junctions may provide the pathway for EDHF in mouse mesenteric artery. J Vasc Res 40: 480–90, 2003. doi: 10.1159/000074549. [ DOI ] [ PubMed ] [ Google Scholar ] 145. Straub AC, Johnstone SR, Heberlein KR, Rizzo MJ, Best AK, Boitano S, Isakson BE. Site-specific connexin phosphorylation is associated with reduced heterocellular communication between smooth muscle and endothelium. J Vasc Res 47: 277–86, 2010. doi: 10.1159/000265562. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 146. Luse MA, Dunaway LS, Nyshadham S, Carvalho A, Sedovy MW, Ruddiman CA, Tessema R, Hirschi K, Johnstone SR, Isakson BE. Endothelial-adipocyte Cx43 Mediated Gap Junctions Can Regulate Adiposity. Function (Oxf) 5, 2024. doi: 10.1093/function/zqae029. [ DOI ] [ Google Scholar ] 147. Siegl D, Koeppen M, Wölfle SE, Pohl U, de Wit C. Myoendothelial coupling is not prominent in arterioles within the mouse cremaster microcirculation in vivo. Circ Res 97: 781–8, 2005. doi: 10.1161/01.RES.0000186193.22438.6c. [ DOI ] [ PubMed ] [ Google Scholar ] 148. Segal SS, Duling BR. Conduction of vasomotor responses in arterioles: a role for cell-to-cell coupling? Am J Physiol 256: H838–45, 1989. doi: 10.1152/ajpheart.1989.256.3.H838. [ DOI ] [ PubMed ] [ Google Scholar ] 149. Biwer LA, Good ME, Hong K, Patel RK, Agrawal N, Looft-Wilson R, Sonkusare SK, Isakson BE. Non-Endoplasmic Reticulum-Based Calr (Calreticulin) Can Coordinate Heterocellular Calcium Signaling and Vascular Function. Arterioscler Thromb Vasc Biol 38: 120–130, 2018. doi: 10.1161/ATVBAHA.117.309886. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 150. Keller TCS, Lechauve C, Keller AS, Broseghini-Filho GB, Butcher JT, Askew Page HR, Islam A, Tan ZY, DeLalio LJ, Brooks S, Sharma P, Hong K, Xu W, Padilha AS, Ruddiman CA, Best AK, Macal E, Kim-Shapiro DB, Christ G, Yan Z, Cortese-Krott MM, Ricart K, Patel R, Bender TP, Sonkusare SK, Weiss MJ, Ackerman H, Columbus L, Isakson BE. Endothelial alpha globin is a nitrite reductase. Nat Commun 13: 6405, 2022. doi: 10.1038/s41467-022-34154-3. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 151. Lamalice L, Le Boeuf F, Huot J. Endothelial cell migration during angiogenesis. Circ Res 100: 782–94, 2007. doi: 10.1161/01.RES.0000259593.07661.1e. [ DOI ] [ PubMed ] [ Google Scholar ] 152. Moccia F, Negri S, Shekha M, Faris P, Guerra G. Endothelial Ca2+ Signaling, Angiogenesis and Vasculogenesis: just What It Takes to Make a Blood Vessel. Int J Mol Sci 20, 2019. doi: 10.3390/ijms20163962. [ DOI ] [ Google Scholar ] 153. Dalal PJ, Muller WA, Sullivan DP. Endothelial Cell Calcium Signaling during Barrier Function and Inflammation. Am J Pathol 190: 535–542, 2020. doi: 10.1016/j.ajpath.2019.11.004. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 154. Dalal PJ, Sullivan DP, Weber EW, Sacks DB, Gunzer M, Grumbach IM, Heller Brown J, Muller WA. Spatiotemporal restriction of endothelial cell calcium signaling is required during leukocyte transmigration. J Exp Med 218, 2021. doi: 10.1084/jem.20192378. [ DOI ] [ Google Scholar ] 155. Bagher P, Segal SS. Regulation of blood flow in the microcirculation: role of conducted vasodilation. Acta Physiol (Oxf) 202: 271–84, 2011. doi: 10.1111/j.1748-1716.2010.02244.x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 156. Paredes RM, Etzler JC, Watts LT, Zheng W, Lechleiter JD. Chemical calcium indicators. Methods 46: 143–51, 2008. doi: 10.1016/j.ymeth.2008.09.025. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 157. Pires PW, Earley S. Neuroprotective effects of TRPA1 channels in the cerebral endothelium following ischemic stroke. Elife 7, 2018. doi: 10.7554/eLife.35316. [ DOI ] [ Google Scholar ] 158. Falcone JC, Kuo L, Meininger GA. Endothelial cell calcium increases during flow-induced dilation in isolated arterioles. Am J Physiol 264: H653–9, 1993. doi: 10.1152/ajpheart.1993.264.2.H653. [ DOI ] [ PubMed ] [ Google Scholar ] 159. Yashiro Y, Duling BR. Participation of intracellular Ca2+ stores in arteriolar conducted responses. Am J Physiol Heart Circ Physiol 285: H65–73, 2003. doi: 10.1152/ajpheart.00662.2002. [ DOI ] [ PubMed ] [ Google Scholar ] 160. Dora KA, Doyle MP, Duling BR. Elevation of intracellular calcium in smooth muscle causes endothelial cell generation of NO in arterioles. Proc Natl Acad Sci U S A 94: 6529–34, 1997. doi: 10.1073/pnas.94.12.6529. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 161. Bagher P, Beleznai T, Kansui Y, Mitchell R, Garland CJ, Dora KA. Low intravascular pressure activates endothelial cell TRPV4 channels, local Ca2+ events, and IKCa channels, reducing arteriolar tone. Proc Natl Acad Sci U S A 109: 18174–9, 2012. doi: 10.1073/pnas.1211946109. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 162. Dora KA, Lin J, Borysova L, Beleznai T, Taggart M, Ascione R, Garland C. Signaling and structures underpinning conducted vasodilation in human and porcine intramyocardial coronary arteries. Front Cardiovasc Med 9: 980628, 2022. doi: 10.3389/fcvm.2022.980628. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 163. Muoboghare MO, Drummond RM, Kennedy C. Characterisation of P2Y2 receptors in human vascular endothelial cells using AR-C118925XX, a competitive and selective P2Y2 antagonist. Br J Pharmacol 176: 2894–2904, 2019. doi: 10.1111/bph.14715. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 164. Nausch LWM, Bonev AD, Heppner TJ, Tallini Y, Kotlikoff MI, Nelson MT. Sympathetic nerve stimulation induces local endothelial Ca2+ signals to oppose vasoconstriction of mouse mesenteric arteries. Am J Physiol Heart Circ Physiol 302: H594–602, 2012. doi: 10.1152/ajpheart.00773.2011. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 165. Castorena-Gonzalez JA, Zawieja SD, Li M, Srinivasan RS, Simon AM, de Wit C, de la Torre R, Martinez-Lemus LA, Hennig GW, Davis MJ. Mechanisms of Connexin-Related Lymphedema. Circ Res 123: 964–985, 2018. doi: 10.1161/CIRCRESAHA.117.312576. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 166. Chen YL, Baker TM, Lee F, Shui B, Lee JC, Tvrdik P, Kotlikoff MI, Sonkusare SK. Calcium Signal Profiles in Vascular Endothelium from Cdh5-GCaMP8 and Cx40-GCaMP2 Mice. J Vasc Res 58: 159–171, 2021. doi: 10.1159/000514210. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 167. Shui B, Lee JC, Reining S, Lee FK, Kotlikoff MI. Optogenetic sensors and effectors: CHROMus-the Cornell Heart Lung Blood Institute Resource for Optogenetic Mouse Signaling. Front Physiol 5: 428, 2014. doi: 10.3389/fphys.2014.00428. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 168. Lee FK, Lee JC, Shui B, Reining S, Jibilian M, Small DM, Jones JS, Allan-Rahill NH, Lamont MR, Rizzo MA, Tajada S, Navedo MF, Santana LF, Nishimura N, Kotlikoff MI. Genetically engineered mice for combinatorial cardiovascular optobiology. Elife 10, 2021. doi: 10.7554/eLife.67858. [ DOI ] [ Google Scholar ] 169. Kotlikoff MI. Genetically encoded Ca2+ indicators: using genetics and molecular design to understand complex physiology. J Physiol 578: 55–67, 2007. doi: 10.1113/jphysiol.2006.120212. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 170. Wang Q, Shui B, Kotlikoff MI, Sondermann H. Structural basis for calcium sensing by GCaMP2. Structure 16: 1817–27, 2008. doi: 10.1016/j.str.2008.10.008. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 171. Tallini YN, Ohkura M, Choi B-R, Ji G, Imoto K, Doran R, Lee J, Plan P, Wilson J, Xin H-B, Sanbe A, Gulick J, Mathai J, Robbins J, Salama G, Nakai J, Kotlikoff MI. Imaging cellular signals in the heart in vivo: Cardiac expression of the high-signal Ca2+ indicator GCaMP2. Proc Natl Acad Sci U S A 103: 4753–8, 2006. doi: 10.1073/pnas.0509378103. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 172. Bagher P, Segal SS. The mouse cremaster muscle preparation for intravital imaging of the microcirculation. J Vis Exp , 2011. doi: 10.3791/2874. [ DOI ] [ Google Scholar ] 173. Bagher P, Davis MJ, Segal SS. Intravital macrozoom imaging and automated analysis of endothelial cell calcium signals coincident with arteriolar dilation in Cx40(BAC) -GCaMP2 transgenic mice. Microcirculation 18: 331–8, 2011. doi: 10.1111/j.1549-8719.2011.00093.x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 174. Bagher P, Davis MJ, Segal SS. Visualizing calcium responses to acetylcholine convection along endothelium of arteriolar networks in Cx40BAC-GCaMP2 transgenic mice. Am J Physiol Heart Circ Physiol 301: H794–802, 2011. doi: 10.1152/ajpheart.00425.2011. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 175. Ye X, Beckett T, Bagher P, Garland CJ, Dora KA. VEGF-A inhibits agonist-mediated Ca2+ responses and activation of IKCa channels in mouse resistance artery endothelial cells. J Physiol 596: 3553–3566, 2018. doi: 10.1113/JP275793. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 176. Mughal A, Hennig GW, Heppner T, Tsoukias NM, Hill-Eubanks D, Nelson MT. Electrocalcium coupling in brain capillaries: Rapidly traveling electrical signals ignite local calcium signals. Proc Natl Acad Sci U S A 121: e2415047121, 2024. doi: 10.1073/pnas.2415047121. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 177. Madisen L, Garner AR, Shimaoka D, Chuong AS, Klapoetke NC, Li L, van der Bourg A, Niino Y, Egolf L, Monetti C, Gu H, Mills M, Cheng A, Tasic B, Nguyen TN, Sunkin SM, Benucci A, Nagy A, Miyawaki A, Helmchen F, Empson RM, Knöpfel T, Boyden ES, Reid RC, Carandini M, Zeng H. Transgenic mice for intersectional targeting of neural sensors and effectors with high specificity and performance. Neuron 85: 942–58, 2015. doi: 10.1016/j.neuron.2015.02.022. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 178. Nelson M, Ledoux J, Taylor M, Bonev A, Hannah R, Solodushko V, Shui B, Tallini Y, Kotlikoff M. Spinning Disk Confocal Microscopy of Calcium Signalling in Blood Vessel Walls. Microsc Anal (Am Ed) 24: 5–8, 2010. [ PMC free article ] [ PubMed ] [ Google Scholar ] 179. Longden TA, Mughal A, Hennig GW, Harraz OF, Shui B, Lee FK, Lee JC, Reining S, Kotlikoff MI, König GM, Kostenis E, Hill-Eubanks D, Nelson MT. Local IP3 receptor-mediated Ca2+ signals compound to direct blood flow in brain capillaries. Sci Adv 7, 2021. doi: 10.1126/sciadv.abh0101. [ DOI ] [ Google Scholar ] 180. Uhlén P. Spectral analysis of calcium oscillations. Sci STKE 2004: pl15, 2004. doi: 10.1126/stke.2582004pl15. [ DOI ] [ PubMed ] [ Google Scholar ] 181. Taylor MS, Francis M. Decoding dynamic Ca(2+) signaling in the vascular endothelium. Front Physiol 5: 447, 2014. doi: 10.3389/fphys.2014.00447. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 182. Ottolini M, Hong K, Sonkusare SK. Calcium signals that determine vascular resistance. Wiley Interdiscip Rev Syst Biol Med 11: e1448, 2019. doi: 10.1002/wsbm.1448. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 183. Moccia F, Brunetti V, Soda T, Berra-Romani R, Scarpellino G. Cracking the Endothelial Calcium (Ca2+) Code: A Matter of Timing and Spacing. Int J Mol Sci 24, 2023. doi: 10.3390/ijms242316765. [ DOI ] [ Google Scholar ] 184. Taylor MS, Francis M, Qian X, Solodushko V. Dynamic Ca(2+) signal modalities in the vascular endothelium. Microcirculation 19: 423–9, 2012. doi: 10.1111/j.1549-8719.2012.00180.x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 185. Thakore P, Earley S. Transient Receptor Potential Channels and Endothelial Cell Calcium Signaling. Compr Physiol 9: 1249–1277, 2019. doi: 10.1002/cphy.c180034. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 186. Bootman M, Niggli E, Berridge M, Lipp P. Imaging the hierarchical Ca2+ signalling system in HeLa cells. J Physiol 499 (Pt 2): 307–14, 1997. doi: 10.1113/jphysiol.1997.sp021928. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 187. Yao Y, Choi J, Parker I. Quantal puffs of intracellular Ca2+ evoked by inositol trisphosphate in Xenopus oocytes. J Physiol 482 (Pt 3): 533–53, 1995. doi: 10.1113/jphysiol.1995.sp020538. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 188. Thomas D, Lipp P, Tovey SC, Berridge MJ, Li W, Tsien RY, Bootman MD. Microscopic properties of elementary Ca2+ release sites in non-excitable cells. Curr Biol 10: 8–15, 2000. doi: 10.1016/s0960-9822(99)00258-4. [ DOI ] [ PubMed ] [ Google Scholar ] 189. Berridge MJ, Lipp P, Bootman MD. The versatility and universality of calcium signalling. Nat Rev Mol Cell Biol 1: 11–21, 2000. doi: 10.1038/35036035. [ DOI ] [ PubMed ] [ Google Scholar ] 190. Saleem H, Tovey SC, Molinski TF, Taylor CW. Interactions of antagonists with subtypes of inositol 1,4,5-trisphosphate (IP3) receptor. Br J Pharmacol 171: 3298–312, 2014. doi: 10.1111/bph.12685. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 191. Clifford PS, Ella SR, Stupica AJ, Nourian Z, Li M, Martinez-Lemus LA, Dora KA, Yang Y, Davis MJ, Pohl U, Meininger GA, Hill MA. Spatial distribution and mechanical function of elastin in resistance arteries: a role in bearing longitudinal stress. Arterioscler Thromb Vasc Biol 31: 2889–96, 2011. doi: 10.1161/ATVBAHA.111.236570. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 192. Pires PW, Sullivan MN, Pritchard HAT, Robinson JJ, Earley S. Unitary TRPV3 channel Ca2+ influx events elicit endothelium-dependent dilation of cerebral parenchymal arterioles. Am J Physiol Heart Circ Physiol 309: H2031–41, 2015. doi: 10.1152/ajpheart.00140.2015. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 193. Sullivan MN, Gonzales AL, Pires PW, Bruhl A, Leo MD, Li W, Oulidi A, Boop FA, Feng Y, Jaggar JH, Welsh DG, Earley S. Localized TRPA1 channel Ca2+ signals stimulated by reactive oxygen species promote cerebral artery dilation. Sci Signal 8: ra2, 2015. doi: 10.1126/scisignal.2005659. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 194. Peters EC, Gee MT, Pawlowski LN, Kath AM, Polk FD, Vance CJ, Sacoman JL, Pires PW. Amyloid-β disrupts unitary calcium entry through endothelial NMDA receptors in mouse cerebral arteries. J Cereb Blood Flow Metab 42: 145–161, 2022. doi: 10.1177/0271678X211039592. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 195. Rahman A, Hughes A, Matchkov V, Nilsson H, Aalkjaer C. Antiphase oscillations of endothelium and smooth muscle [Ca2+]i in vasomotion of rat mesenteric small arteries. Cell Calcium 42: 536–47, 2007. doi: 10.1016/j.ceca.2007.01.007. [ DOI ] [ PubMed ] [ Google Scholar ] 196. Francis M, Qian X, Charbel C, Ledoux J, Parker JC, Taylor MS. Automated region of interest analysis of dynamic Ca 2 + signals in image sequences. Am J Physiol Cell Physiol 303: C236–43, 2012. doi: 10.1152/ajpcell.00016.2012. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 197. Taylor MS, Choi C-S, Bayazid L, Glosemeyer KE, Baker CCP, Weber DS. Changes in vascular reactivity and endothelial Ca2+ dynamics with chronic low flow. Microcirculation 24, 2017. doi: 10.1111/micc.12354. [ DOI ] [ Google Scholar ] 198. Wilson C, Saunter CD, Girkin JM, McCarron JG. Pressure-dependent regulation of Ca2+ signalling in the vascular endothelium. J Physiol 593: 5231–53, 2015. doi: 10.1113/JP271157. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 199. Breslin JW. Edema and lymphatic clearance: molecular mechanisms and ongoing challenges. Clin Sci (Lond) 137: 1451–1476, 2023. doi: 10.1042/CS20220314. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 200. Durán WN, Sánchez FA, Breslin JW. Microcirculatory Exchange Function. In: Handbook of Physiology: Microcirculation. Elsevier/Academic Press, 2008. [ Google Scholar ] 201. Yuan SY, Rigir RR. Regulation of Endothelial Barrier Function. In: Regulation of Endothelial Barrier Function. 2010. [ Google Scholar ] 202. Villalba N, Ma Y, Gahan SA, Joly-Amado A, Spence S, Yang X, Nash KR, Yuan SY. Lung infection by Pseudomonas aeruginosa induces neuroinflammation and blood-brain barrier dysfunction in mice. J Neuroinflammation 20: 127, 2023. doi: 10.1186/s12974-023-02817-7. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 203. Villalba N, Baby S, Cha BJ, Yuan SY. Site-specific opening of the blood-brain barrier by extracellular histones. J Neuroinflammation 17: 281, 2020. doi: 10.1186/s12974-020-01950-x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 204. Saria A, Lundberg JM. Evans blue fluorescence: quantitative and morphological evaluation of vascular permeability in animal tissues. J Neurosci Methods 8: 41–9, 1983. doi: 10.1016/0165-0270(83)90050-x. [ DOI ] [ PubMed ] [ Google Scholar ] 205. MILES AA, MILES EM. Vascular reactions to histamine, histamine-liberator and leukotaxine in the skin of guinea-pigs. J Physiol 118: 228–57, 1952. doi: 10.1113/jphysiol.1952.sp004789. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 206. Svensjö E, Arfors KE, Arturson G, Rutili G. The hamster cheek pouch preparation as a model for studies of macromolecular permeability of the microvasculature. Ups J Med Sci 83: 71–9, 1978. doi: 10.3109/03009737809179115. [ DOI ] [ PubMed ] [ Google Scholar ] 207. Gawlowski DM, Ritter AB, Durán WN. Reproducibility of microvascular permeability responses to successive topical applications of bradykinin in the hamster cheek pouch. Microvasc Res 24: 354–63, 1982. doi: 10.1016/0026-2862(82)90022-x. [ DOI ] [ PubMed ] [ Google Scholar ] 208. Bekker AY, Ritter AB, Durán WN. Analysis of microvascular permeability to macromolecules by video-image digital processing. Microvasc Res 38: 200–16, 1989. doi: 10.1016/0026-2862(89)90028-9. [ DOI ] [ PubMed ] [ Google Scholar ] 209. Doggett TM, Tur JJ, Alves NG, Yuan SY, Tipparaju SM, Breslin JW. Assessment of Cardiovascular Function and Microvascular Permeability in a Conscious Rat Model of Alcohol Intoxication Combined with Hemorrhagic Shock and Resuscitation. Methods Mol Biol 1717: 61–81, 2018. doi: 10.1007/978-1-4939-7526-6_6. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 210. Ramírez MM, Quardt SM, Kim D, Oshiro H, Minnicozzi M, Durán WN. Platelet activating factor modulates microvascular permeability through nitric oxide synthesis. Microvasc Res 50: 223–34, 1995. doi: 10.1006/mvre.1995.1055. [ DOI ] [ PubMed ] [ Google Scholar ] 211. Kim D, Armenante PM, Durán WN. Transient analysis of macromolecular transport across microvascular wall and into interstitium. Am J Physiol 265: H993–9, 1993. doi: 10.1152/ajpheart.1993.265.3.H993. [ DOI ] [ PubMed ] [ Google Scholar ] 212. Alves NG, Trujillo AN, Breslin JW, Yuan SY. Sphingosine-1-Phosphate Reduces Hemorrhagic Shock and Resuscitation-Induced Microvascular Leakage by Protecting Endothelial Mitochondrial Integrity. Shock 52: 423–433, 2019. doi: 10.1097/SHK.0000000000001280. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 213. Breslin JW, Wu MH, Guo M, Reynoso R, Yuan SY. Toll-like receptor 4 contributes to microvascular inflammation and barrier dysfunction in thermal injury. Shock 29: 349–55, 2008. doi: 10.1097/shk.0b013e3181454975. [ DOI ] [ PubMed ] [ Google Scholar ] 214. Reynoso R, Perrin RM, Breslin JW, Daines DA, Watson KD, Watterson DM, Wu MH, Yuan S. A role for long chain myosin light chain kinase (MLCK-210) in microvascular hyperpermeability during severe burns. Shock 28: 589–95, 2007. doi: 10.1097/SHK.0b013e31804d415f. [ DOI ] [ PubMed ] [ Google Scholar ] 215. Tharakan B, Whaley JG, Hunter FA, Smythe WR, Childs EW. (-)-Deprenyl inhibits vascular hyperpermeability after hemorrhagic shock. Shock 33: 56–63, 2010. doi: 10.1097/SHK.0b013e3181a7fb7c. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 216. Huxley VH, Curry FE, Adamson RH. Quantitative fluorescence microscopy on single capillaries: alpha-lactalbumin transport. Am J Physiol 252: H188–97, 1987. doi: 10.1152/ajpheart.1987.252.1.H188. [ DOI ] [ PubMed ] [ Google Scholar ] 217. Sarelius IH, Kuebel JM, Wang J, Huxley VH. Macromolecule permeability of in situ and excised rodent skeletal muscle arterioles and venules. Am J Physiol Heart Circ Physiol 290: H474–80, 2006. doi: 10.1152/ajpheart.00655.2005. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 218. Yuan Y, Chilian WM, Granger HJ, Zawieja DC. Permeability to albumin in isolated coronary venules. Am J Physiol 265: H543–52, 1993. doi: 10.1152/ajpheart.1993.265.2.H543. [ DOI ] [ PubMed ] [ Google Scholar ] 219. Yuan Y, Granger HJ, Zawieja DC, Chilian WM. Flow modulates coronary venular permeability by a nitric oxide-related mechanism. Am J Physiol 263: H641–6, 1992. doi: 10.1152/ajpheart.1992.263.2.H641. [ DOI ] [ PubMed ] [ Google Scholar ] 220. Yuan Y, Granger HJ, Zawieja DC, DeFily DV, Chilian WM. Histamine increases venular permeability via a phospholipase C-NO synthase-guanylate cyclase cascade. Am J Physiol 264: H1734–9, 1993. doi: 10.1152/ajpheart.1993.264.5.H1734. [ DOI ] [ PubMed ] [ Google Scholar ] 221. Yuan Y, Huang Q, Wu HM. Myosin light chain phosphorylation: modulation of basal and agonist-stimulated venular permeability. Am J Physiol 272: H1437–43, 1997. doi: 10.1152/ajpheart.1997.272.3.H1437. [ DOI ] [ PubMed ] [ Google Scholar ] 222. Michel CC. Fluid movements through capillary walls. In: Handbook of Physiology: The Cardiovascular System Microcirculation. 1984, p. 375–409. [ Google Scholar ] 223. Breslin JW, Yuan SY. Involvement of RhoA and Rho kinase in neutrophil-stimulated endothelial hyperpermeability. Am J Physiol Heart Circ Physiol 286: H1057–62, 2004. doi: 10.1152/ajpheart.00841.2003. [ DOI ] [ PubMed ] [ Google Scholar ] 224. Breslin JW, Daines DA, Doggett TM, Kurtz KH, Souza-Smith FM, Zhang XE, Wu MH, Yuan SY. Rnd3 as a Novel Target to Ameliorate Microvascular Leakage. J Am Heart Assoc 5: e003336, 2016. doi: 10.1161/JAHA.116.003336. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 225. Yuan SY, Wu MH, Ustinova EE, Guo M, Tinsley JH, De Lanerolle P, Xu W. Myosin light chain phosphorylation in neutrophil-stimulated coronary microvascular leakage. Circ Res 90: 1214–21, 2002. doi: 10.1161/01.res.0000020402.73609.f1. [ DOI ] [ PubMed ] [ Google Scholar ] 226. Yuan SY. Signal transduction pathways in enhanced microvascular permeability. Microcirculation 7: 395–403, 2000. [ PubMed ] [ Google Scholar ] 227. Tinsley JH, Breslin JW, Teasdale NR, Yuan SY. PKC-dependent, burn-induced adherens junction reorganization and barrier dysfunction in pulmonary microvascular endothelial cells. Am J Physiol Lung Cell Mol Physiol 289: L217–23, 2005. doi: 10.1152/ajplung.00248.2004. [ DOI ] [ PubMed ] [ Google Scholar ] 228. Breslin JW, Pappas PJ, Cerveira JJ, Hobson RW, Durán WN. VEGF increases endothelial permeability by separate signaling pathways involving ERK-1/2 and nitric oxide. Am J Physiol Heart Circ Physiol 284: H92–H100, 2003. doi: 10.1152/ajpheart.00330.2002. [ DOI ] [ PubMed ] [ Google Scholar ] 229. Breslin JW, Sun H, Xu W, Rodarte C, Moy AB, Wu MH, Yuan SY. Involvement of ROCK-mediated endothelial tension development in neutrophil-stimulated microvascular leakage. Am J Physiol Heart Circ Physiol 290: H741–50, 2006. doi: 10.1152/ajpheart.00238.2005. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 230. Alves NG, Yuan SY, Breslin JW. Sphingosine-1-phosphate protects against brain microvascular endothelial junctional protein disorganization and barrier dysfunction caused by alcohol. Microcirculation 26: e12506, 2019. doi: 10.1111/micc.12506. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 231. Heng BC, Bezerra PP, Preiser PR, Law SKA, Xia Y, Boey F, Venkatraman SS. Effect of cell-seeding density on the proliferation and gene expression profile of human umbilical vein endothelial cells within ex vivo culture. Cytotherapy 13: 606–17, 2011. doi: 10.3109/14653249.2010.542455. [ DOI ] [ PubMed ] [ Google Scholar ] 232. Rigor RR, Beard RS, Litovka OP, Yuan SY. Interleukin-1β-induced barrier dysfunction is signaled through PKC-θ in human brain microvascular endothelium. Am J Physiol Cell Physiol 302: C1513–22, 2012. doi: 10.1152/ajpcell.00371.2011. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 233. Beard RS, Hoettels BA, Meegan JE, Wertz TS, Cha BJ, Yang X, Oxford JT, Wu MH, Yuan SY. AKT2 maintains brain endothelial claudin-5 expression and selective activation of IR/AKT2/FOXO1-signaling reverses barrier dysfunction. J Cereb Blood Flow Metab 40: 374–391, 2020. doi: 10.1177/0271678X18817512. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 234. Beard RS, Haines RJ, Wu KY, Reynolds JJ, Davis SM, Elliott JE, Malinin NL, Chatterjee V, Cha BJ, Wu MH, Yuan SY. Non-muscle Mlck is required for β-catenin- and FoxO1-dependent downregulation of Cldn5 in IL-1β-mediated barrier dysfunction in brain endothelial cells. J Cell Sci 127: 1840–53, 2014. doi: 10.1242/jcs.144550. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 235. Robinson BD, Shaji CA, Lomas A, Tharakan B. Measurement of Microvascular Endothelial Barrier Dysfunction and Hyperpermeability In Vitro. Methods Mol Biol 1717: 237–242, 2018. doi: 10.1007/978-1-4939-7526-6_19. [ DOI ] [ PubMed ] [ Google Scholar ] 236. Bingaman S, Huxley VH, Rumbaut RE. Fluorescent dyes modify properties of proteins used in microvascular research. Microcirculation 10: 221–31, 2003. doi: 10.1038/sj.mn.7800186. [ DOI ] [ PubMed ] [ Google Scholar ] 237. Motawe ZY, Farsaei F, Abdelmaboud SS, Cuevas J, Breslin JW. Sigma-1 receptor activation-induced glycolytic ATP production and endothelial barrier enhancement. Microcirculation 27: e12620, 2020. doi: 10.1111/micc.12620. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 238. Breslin JW, Yuan SY. Determination of Solute Permeability of Microvascular Endothelial Cell Monolayers In Vitro. Methods Mol Biol 2711: 1–12, 2024. doi: 10.1007/978-1-0716-3429-5_1. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 239. Albelda SM, Sampson PM, Haselton FR, McNiff JM, Mueller SN, Williams SK, Fishman AP, Levine EM. Permeability characteristics of cultured endothelial cell monolayers. J Appl Physiol (1985) 64: 308–22, 1988. doi: 10.1152/jappl.1988.64.1.308. [ DOI ] [ PubMed ] [ Google Scholar ] 240. Shasby DM, Shasby SS. Effects of calcium on transendothelial albumin transfer and electrical resistance. J Appl Physiol (1985) 60: 71–9, 1986. doi: 10.1152/jappl.1986.60.1.71. [ DOI ] [ PubMed ] [ Google Scholar ] 241. Navab M, Hough GP, Berliner JA, Frank JA, Fogelman AM, Haberland ME, Edwards PA. Rabbit beta-migrating very low density lipoprotein increases endothelial macromolecular transport without altering electrical resistance. J Clin Invest 78: 389–97, 1986. doi: 10.1172/JCI112589. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 242. Territo M, Berliner JA, Fogelman AM. Effect of monocyte migration on low density lipoprotein transport across aortic endothelial cell monolayers. J Clin Invest 74: 2279–84, 1984. doi: 10.1172/JCI111655. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 243. Furie MB, Cramer EB, Naprstek BL, Silverstein SC. Cultured endothelial cell monolayers that restrict the transendothelial passage of macromolecules and electrical current. J Cell Biol 98: 1033–41, 1984. doi: 10.1083/jcb.98.3.1033. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 244. Raub TJ, Kuentzel SL, Sawada GA. Permeability of bovine brain microvessel endothelial cells in vitro: barrier tightening by a factor released from astroglioma cells. Exp Cell Res 199: 330–40, 1992. doi: 10.1016/0014-4827(92)90442-b. [ DOI ] [ PubMed ] [ Google Scholar ] 245. Abbruscato TJ, Davis TP. Combination of hypoxia/aglycemia compromises in vitro blood-brain barrier integrity. J Pharmacol Exp Ther 289: 668–75, 1999. [ PubMed ] [ Google Scholar ] 246. Tilling T, Korte D, Hoheisel D, Galla HJ. Basement membrane proteins influence brain capillary endothelial barrier function in vitro. J Neurochem 71: 1151–7, 1998. doi: 10.1046/j.1471-4159.1998.71031151.x. [ DOI ] [ PubMed ] [ Google Scholar ] 247. Giaever I, Keese CR. Micromotion of mammalian cells measured electrically. Proc Natl Acad Sci U S A 88: 7896–900, 1991. doi: 10.1073/pnas.88.17.7896. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 248. Giaever I, Keese CR. A morphological biosensor for mammalian cells. Nature 366: 591–2, 1993. doi: 10.1038/366591a0. [ DOI ] [ PubMed ] [ Google Scholar ] 249. Moy AB, Van Engelenhoven J, Bodmer J, Kamath J, Keese C, Giaever I, Shasby S, Shasby DM. Histamine and thrombin modulate endothelial focal adhesion through centripetal and centrifugal forces. J Clin Invest 97: 1020–7, 1996. doi: 10.1172/JCI118493. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 250. Moy AB, Blackwell K, Wang N, Haxhinasto K, Kasiske MK, Bodmer J, Reyes G, English A. Phorbol ester-mediated pulmonary artery endothelial barrier dysfunction through regulation of actin cytoskeletal mechanics. Am J Physiol Lung Cell Mol Physiol 287: L153–67, 2004. doi: 10.1152/ajplung.00292.2003. [ DOI ] [ PubMed ] [ Google Scholar ] 251. Moy AB, Blackwell K, Kamath A. Differential effects of histamine and thrombin on endothelial barrier function through actin-myosin tension. Am J Physiol Heart Circ Physiol 282: H21–9, 2002. doi: 10.1152/ajpheart.2002.282.1.H21. [ DOI ] [ PubMed ] [ Google Scholar ] 252. Miller NE, Michel CC, Nanjee MN, Olszewski WL, Miller IP, Hazell M, Olivecrona G, Sutton P, Humphreys SM, Frayn KN. Secretion of adipokines by human adipose tissue in vivo: partitioning between capillary and lymphatic transport. Am J Physiol Endocrinol Metab 301: E659–67, 2011. doi: 10.1152/ajpendo.00058.2011. [ DOI ] [ PubMed ] [ Google Scholar ] 253. Kohan AB, Yoder SM, Tso P. Using the lymphatics to study nutrient absorption and the secretion of gastrointestinal hormones. Physiol Behav 105: 82–8, 2011. doi: 10.1016/j.physbeh.2011.04.056. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 254. Michel CC, Woodcock TE, Curry F- RE. Understanding and extending the Starling principle. Acta Anaesthesiol Scand 64: 1032–1037, 2020. doi: 10.1111/aas.13603. [ DOI ] [ PubMed ] [ Google Scholar ] 255. Tso P, Pitts V, Granger DN. Role of lymph flow in intestinal chylomicron transport. Am J Physiol 249: G21–8, 1985. doi: 10.1152/ajpgi.1985.249.1.G21. [ DOI ] [ PubMed ] [ Google Scholar ] 256. Dongaonkar RM, Nguyen TL, Quick CM, Hardy J, Laine GA, Wilson E, Stewart RH. Adaptation of mesenteric lymphatic vessels to prolonged changes in transmural pressure. Am J Physiol Heart Circ Physiol 305: H203–10, 2013. doi: 10.1152/ajpheart.00677.2012. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 257. Gashev AA, Davis MJ, Delp MD, Zawieja DC. Regional variations of contractile activity in isolated rat lymphatics. Microcirculation 11: 477–92, 2004. doi: 10.1080/10739680490476033. [ DOI ] [ PubMed ] [ Google Scholar ] 258. Quick CM, Venugopal AM, Gashev AA, Zawieja DC, Stewart RH. Intrinsic pump-conduit behavior of lymphangions. Am J Physiol Regul Integr Comp Physiol 292: R1510–8, 2007. doi: 10.1152/ajpregu.00258.2006. [ DOI ] [ PubMed ] [ Google Scholar ] 259. Baeyens N, Nicoli S, Coon BG, Ross TD, Van den Dries K, Han J, Lauridsen HM, Mejean CO, Eichmann A, Thomas J-L, Humphrey JD, Schwartz MA. Vascular remodeling is governed by a VEGFR3-dependent fluid shear stress set point. Elife 4, 2015. doi: 10.7554/eLife.04645. [ DOI ] [ Google Scholar ] 260. Gashev AA, Davis MJ, Zawieja DC. Inhibition of the active lymph pump by flow in rat mesenteric lymphatics and thoracic duct. J Physiol 540: 1023–37, 2002. doi: 10.1113/jphysiol.2001.016642. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 261. Zawieja SD, Castorena-Gonzalez JA, Scallan JP, Davis MJ. Differences in L-type Ca2+ channel activity partially underlie the regional dichotomy in pumping behavior by murine peripheral and visceral lymphatic vessels. Am J Physiol Heart Circ Physiol 314: H991–H1010, 2018. doi: 10.1152/ajpheart.00499.2017. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 262. Van Helden DF. Pacemaker potentials in lymphatic smooth muscle of the guinea-pig mesentery. J Physiol 471: 465–79, 1993. doi: 10.1113/jphysiol.1993.sp019910. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 263. Barankay T, Baumgärtl H, Lübbers DW, Seidl E. Oxygen pressure in small lymphatics. Pflugers Arch 366: 53–9, 1976. doi: 10.1007/BF02486560. [ DOI ] [ PubMed ] [ Google Scholar ] 264. Hangai-Hoger N, Tsai AG, Cabrales P, Intaglietta M. Terminal lymphatics: the potential “lethal corner” in the distribution of tissue pO2. Lymphat Res Biol 5: 159–68, 2007. doi: 10.1089/lrb.2007.5303. [ DOI ] [ PubMed ] [ Google Scholar ] 265. Witte CL, Clauss RH, Dumont AE. Respiratory gas tensions of thoracic duct lymph: an index of gas exchange in splanchnic tissues. Ann Surg 166: 254–62, 1967. doi: 10.1097/00000658-196708000-00013. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 266. Ohhashi T, Kawai Y. Proposed new lymphology combined with lymphatic physiology, innate immunology, and oncology. J Physiol Sci 65: 51–66, 2015. doi: 10.1007/s12576-014-0343-6. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 267. Briggs Boedtkjer D, Rumessen J, Baandrup U, Skov Mikkelsen M, Telinius N, Pilegaard H, Aalkjaer C, Hjortdal V. Identification of interstitial Cajal-like cells in the human thoracic duct. Cells Tissues Organs 197: 145–58, 2013. doi: 10.1159/000342437. [ DOI ] [ PubMed ] [ Google Scholar ] 268. Telinius N, Drewsen N, Pilegaard H, Kold-Petersen H, de Leval M, Aalkjaer C, Hjortdal V, Boedtkjer DB. Human thoracic duct in vitro: diameter-tension properties, spontaneous and evoked contractile activity. Am J Physiol Heart Circ Physiol 299: H811–8, 2010. doi: 10.1152/ajpheart.01089.2009. [ DOI ] [ PubMed ] [ Google Scholar ] 269. Telinius N, Kim S, Pilegaard H, Pahle E, Nielsen J, Hjortdal V, Aalkjaer C, Boedtkjer DB. The contribution of K(+) channels to human thoracic duct contractility. Am J Physiol Heart Circ Physiol 307: H33–43, 2014. doi: 10.1152/ajpheart.00921.2013. [ DOI ] [ PubMed ] [ Google Scholar ] 270. Telinius N, Majgaard J, Kim S, Katballe N, Pahle E, Nielsen J, Hjortdal V, Aalkjaer C, Boedtkjer DB. Voltage-gated sodium channels contribute to action potentials and spontaneous contractility in isolated human lymphatic vessels. J Physiol 593: 3109–22, 2015. doi: 10.1113/JP270166. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 271. Bixel MG, Sivaraj KK, Timmen M, Mohanakrishnan V, Aravamudhan A, Adams S, Koh B-I, Jeong H-W, Kruse K, Stange R, Adams RH. Angiogenesis is uncoupled from osteogenesis during calvarial bone regeneration. Nat Commun 15: 4575, 2024. doi: 10.1038/s41467-024-48579-5. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 272. Boedtkjer E. Acid-base regulation and sensing: Accelerators and brakes in metabolic regulation of cerebrovascular tone. J Cereb Blood Flow Metab 38: 588–602, 2018. doi: 10.1177/0271678X17733868. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 273. Andreasen F, Christensen JH, Poulsen B. A method of oxygen supply to a low volume tissue bath containing a protein solution. J Pharmacol Methods 17: 277–81, 1987. doi: 10.1016/0160-5402(87)90058-1. [ DOI ] [ PubMed ] [ Google Scholar ] 274. Scallan JP, Davis MJ. Genetic removal of basal nitric oxide enhances contractile activity in isolated murine collecting lymphatic vessels. J Physiol 591: 2139–56, 2013. doi: 10.1113/jphysiol.2012.250662. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 275. Scallan JP, Huxley VH. In vivo determination of collecting lymphatic vessel permeability to albumin: a role for lymphatics in exchange. J Physiol 588: 243–54, 2010. doi: 10.1113/jphysiol.2009.179622. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 276. Zawieja SD, Pea GA, Broyhill SE, Patro A, Bromert KH, Li M, Norton CE, Castorena-Gonzalez JA, Hancock EJ, Bertram CD, Davis MJ. IP3R1 underlies diastolic ANO1 activation and pressure-dependent chronotropy in lymphatic collecting vessels. J Gen Physiol 155, 2023. doi: 10.1085/jgp.202313358. [ DOI ] [ Google Scholar ] 277. Davis MJ, Bertram CD. Control of lymphatic pacemaking and pumping by mechanobiological signals. J Physiol 603: 3307–3327, 2025. doi: 10.1113/JP288477. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 278. DuToit J, Brothers P, Stephens M, Keane K, de Jesus FN, Roizes S, von der Weid P-Y. Flow-dependent regulation of rat mesenteric lymphatic vessel contractile response requires activation of endothelial TRPV4 channels. Microcirculation 31: e12839, 2024. doi: 10.1111/micc.12839. [ DOI ] [ PubMed ] [ Google Scholar ] 279. Aldecoa C, Llau JV, Nuvials X, Artigas A. Role of albumin in the preservation of endothelial glycocalyx integrity and the microcirculation: a review. Ann Intensive Care 10: 85, 2020. doi: 10.1186/s13613-020-00697-1. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 280. Schulz ME, Akerstrom VL, Song K, Broyhill SE, Li M, Lambert MD, Goldberg TB, Kataru RP, Shin J, Braun SE, Norton CE, Czepielewski RS, Mehrara BJ, Domeier TL, Zawieja SD, Castorena-Gonzalez JA. Regulation of Collecting Lymphatic Vessel Contractile Function by TRPV4 Channels. Arterioscler Thromb Vasc Biol 45: e412–e436, 2025. doi: 10.1161/ATVBAHA.124.322100. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 281. Williamson IM. Some responses of bovine mesenteric arteries, veins and lymphatics. J Physiol 202: 112P+, 1969. [ Google Scholar ] 282. Mawhinney HJ, Roddie IC. Spontaneous activity in isolated bovine mesenteric lymphatics. J Physiol 229: 339–48, 1973. doi: 10.1113/jphysiol.1973.sp010141. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 283. McHale NG, Roddie IC. Pumping activity in isolated segments of bovine mesenteric lymphatics. J Physiol 244: 70P–72P, 1975. [ Google Scholar ] 284. Ohhashi T, Kawai Y, Azuma T. The response of lymphatic smooth muscles to vasoactive substances. Pflugers Arch 375: 183–8, 1978. doi: 10.1007/BF00584242. [ DOI ] [ PubMed ] [ Google Scholar ] 285. Johnston MG, Feuer C. Suppression of lymphatic vessel contractility with inhibitors of arachidonic acid metabolism. J Pharmacol Exp Ther 226: 603–7, 1983. [ PubMed ] [ Google Scholar ] 286. Johnston MG, Kanalec A, Gordon JL. Effects of arachidonic acid and its cyclo-oxygenase and lipoxygenase products on lymphatic vessel contractility in vitro. Prostaglandins 25: 85–98, 1983. doi: 10.1016/0090-6980(83)90138-7. [ DOI ] [ PubMed ] [ Google Scholar ] 287. Ono N, Mizuno R, Nojiri H, Ohhashi T. Development of an experimental apparatus for investigating lymphatic pumping activity of murine mesentery in vivo. Jpn J Physiol 50: 25–31, 2000. doi: 10.2170/jjphysiol.50.25. [ DOI ] [ PubMed ] [ Google Scholar ] 288. Liao S, Cheng G, Conner DA, Huang Y, Kucherlapati RS, Munn LL, Ruddle NH, Jain RK, Fukumura D, Padera TP. Impaired lymphatic contraction associated with immunosuppression. Proc Natl Acad Sci U S A 108: 18784–9, 2011. doi: 10.1073/pnas.1116152108. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 289. Telinius N, Drewsen N, Pilegaard H, Kold-Petersen H, de Leval M, Aalkjaer C, Hjortdal V, Boedtkjer DB. Human thoracic duct in vitro: diameter-tension properties, spontaneous and evoked contractile activity. American Journal of Physiology-Heart and Circulatory Physiology 299: H811–H818, 2010. doi: 10.1152/ajpheart.01089.2009. [ DOI ] [ PubMed ] [ Google Scholar ] 290. Moeller AL, Hjortdal VE, Boedtkjer DMB, Boedtkjer E. Acidosis inhibits rhythmic contractions of human thoracic ducts. Physiol Rep 7: e14074, 2019. doi: 10.14814/phy2.14074. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 291. Zhao J, van Helden DF. ET-1-associated vasomotion and vasospasm in lymphatic vessels of the guinea-pig mesentery. Br J Pharmacol 140: 1399–413, 2003. doi: 10.1038/sj.bjp.0705573. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 292. Choi I, Chung HK, Ramu S, Lee HN, Kim KE, Lee S, Yoo J, Choi D, Lee YS, Aguilar B, Hong Y-K. Visualization of lymphatic vessels by Prox1-promoter directed GFP reporter in a bacterial artificial chromosome-based transgenic mouse. Blood 117: 362–5, 2011. doi: 10.1182/blood-2010-07-298562. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 293. Hägerling R, Pollmann C, Kremer L, Andresen V, Kiefer F. Intravital two-photon microscopy of lymphatic vessel development and function using a transgenic Prox1 promoter-directed mOrange2 reporter mouse. Biochem Soc Trans 39: 1674–81, 2011. doi: 10.1042/BST20110722. [ DOI ] [ PubMed ] [ Google Scholar ] 294. Martínez-Corral I, Olmeda D, Diéguez-Hurtado R, Tammela T, Alitalo K, Ortega S. In vivo imaging of lymphatic vessels in development, wound healing, inflammation, and tumor metastasis. Proc Natl Acad Sci U S A 109: 6223–8, 2012. doi: 10.1073/pnas.1115542109. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 295. Bianchi R, Teijeira A, Proulx ST, Christiansen AJ, Seidel CD, Rülicke T, Mäkinen T, Hägerling R, Halin C, Detmar M. A transgenic Prox1-Cre-tdTomato reporter mouse for lymphatic vessel research. PLoS One 10: e0122976, 2015. doi: 10.1371/journal.pone.0122976. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 296. Jung E, Gardner D, Choi D, Park E, Jin Seong Y, Yang S, Castorena-Gonzalez J, Louveau A, Zhou Z, Lee GK, Perrault DP, Lee S, Johnson M, Daghlian G, Lee M, Jin Hong Y, Kato Y, Kipnis J, Davis MJ, Wong AK, Hong Y-K. Development and Characterization of A Novel Prox1-EGFP Lymphatic and Schlemm’s Canal Reporter Rat. Sci Rep 7: 5577, 2017. doi: 10.1038/s41598-017-06031-3. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 297. Razavi MS, Leonard-Duke J, Hardie B, Dixon JB, Gleason RL. Axial stretch regulates rat tail collecting lymphatic vessel contractions. Sci Rep 10: 5918, 2020. doi: 10.1038/s41598-020-62799-x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 298. von der Weid PY, Crowe MJ, Van Helden DF. Endothelium-dependent modulation of pacemaking in lymphatic vessels of the guinea-pig mesentery. J Physiol 493 (Pt 2): 563–75, 1996. doi: 10.1113/jphysiol.1996.sp021404. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 299. Nizamutdinova IT, Maejima D, Nagai T, Bridenbaugh E, Thangaswamy S, Chatterjee V, Meininger CJ, Gashev AA. Involvement of histamine in endothelium-dependent relaxation of mesenteric lymphatic vessels. Microcirculation 21: 640–8, 2014. doi: 10.1111/micc.12143. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 300. Ohhashi T, Takahashi N. Acetylcholine-induced release of endothelium-derived relaxing factor from lymphatic endothelial cells. Am J Physiol 260: H1172–8, 1991. doi: 10.1152/ajpheart.1991.260.4.H1172. [ DOI ] [ PubMed ] [ Google Scholar ] 301. Scallan JP, Jannaway M. Lymphatic Vascular Permeability. Cold Spring Harb Perspect Med 12, 2022. doi: 10.1101/cshperspect.a041274. [ DOI ] [ Google Scholar ] 302. Sabine A, Davis MJ, Bovay E, Petrova TV. Characterization of Mouse Mesenteric Lymphatic Valve Structure and Function. Methods Mol Biol 1846: 97–129, 2018. doi: 10.1007/978-1-4939-8712-2_7. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 303. Davis MJ, Castorena-Gonzalez JA, Li M, Zawieja SD, Simon AM, Geng X, Srinivasan RS. Connexin-45 is expressed in mouse lymphatic endothelium and required for lymphatic valve function. JCI Insight 9, 2024. doi: 10.1172/jci.insight.169931. [ DOI ] [ Google Scholar ] 304. Davis MJ, Zawieja SD, Yang Y. Developmental progression of lymphatic valve morphology and function. Front Cell Dev Biol 12: 1331291, 2024. doi: 10.3389/fcell.2024.1331291. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 305. Lapinski PE, Lubeck BA, Chen D, Doosti A, Zawieja SD, Davis MJ, King PD. RASA1 regulates the function of lymphatic vessel valves in mice. J Clin Invest 127: 2569–2585, 2017. doi: 10.1172/JCI89607. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 306. Davis MJ, Rahbar E, Gashev AA, Zawieja DC, Moore JE. Determinants of valve gating in collecting lymphatic vessels from rat mesentery. Am J Physiol Heart Circ Physiol 301: H48–60, 2011. doi: 10.1152/ajpheart.00133.2011. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 307. Scallan JP, Davis MJ, Huxley VH. Permeability and contractile responses of collecting lymphatic vessels elicited by atrial and brain natriuretic peptides. J Physiol 591: 5071–81, 2013. doi: 10.1113/jphysiol.2013.260042. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 308. Scallan JP, Hill MA, Davis MJ. Lymphatic vascular integrity is disrupted in type 2 diabetes due to impaired nitric oxide signalling. Cardiovasc Res 107: 89–97, 2015. doi: 10.1093/cvr/cvv117. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 309. Jannaway M, Scallan JP. Lymphatic Vascular Permeability Determined from Direct Measurements of Solute Flux. Methods Mol Biol 2711: 21–37, 2024. doi: 10.1007/978-1-0716-3429-5_3. [ DOI ] [ PubMed ] [ Google Scholar ] 310. Chiu J-J, Chien S Effects of disturbed flow on vascular endothelium: pathophysiological basis and clinical perspectives. Physiol Rev 91: 327–87, 2011. doi: 10.1152/physrev.00047.2009. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 311. Davis MJ, Earley S, Li Y-S, Chien S. Vascular mechanotransduction. Physiol Rev 103: 1247–1421, 2023. doi: 10.1152/physrev.00053.2021. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 312. Earley S, Brayden JE. Transient receptor potential channels in the vasculature. Physiol Rev 95: 645–90, 2015. doi: 10.1152/physrev.00026.2014. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 313. Garcia DCG, Longden TA. Ion channels in capillary endothelium. Curr Top Membr 85: 261–300, 2020. doi: 10.1016/bs.ctm.2020.01.005. [ DOI ] [ PubMed ] [ Google Scholar ] 314. Jackson WF. Endothelial Ion Channels and Cell-Cell Communication in the Microcirculation. Front Physiol 13: 805149, 2022. doi: 10.3389/fphys.2022.805149. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 315. Metwally E, Sanchez Solano A, Lavanderos B, Yamasaki E, Thakore P, McClenaghan C, Rios N, Radi R, Feng Earley Y, Nichols CG, Earley S. Mitochondrial Ca2+-coupled generation of reactive oxygen species, peroxynitrite formation, and endothelial dysfunction in Cantú syndrome. JCI Insight 9, 2024. doi: 10.1172/jci.insight.176212. [ DOI ] [ Google Scholar ] 316. Sobey CG. Potassium channel function in vascular disease. Arterioscler Thromb Vasc Biol 21: 28–38, 2001. doi: 10.1161/01.atv.21.1.28. [ DOI ] [ PubMed ] [ Google Scholar ] 317. Nishijima Y, Zheng X, Lund H, Suzuki M, Mattson DL, Zhang DX. Characterization of blood pressure and endothelial function in TRPV4-deficient mice with l-NAME- and angiotensin II-induced hypertension. Physiol Rep 2: e00199, 2014. doi: 10.1002/phy2.199. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 318. Paravicini TM, Yogi A, Mazur A, Touyz RM. Dysregulation of vascular TRPM7 and annexin-1 is associated with endothelial dysfunction in inherited hypomagnesemia. Hypertension 53: 423–9, 2009. doi: 10.1161/HYPERTENSIONAHA.108.124651. [ DOI ] [ PubMed ] [ Google Scholar ] 319. Lim XR, Willemse L, Harraz OF. Amyloid beta Aβ1-40 activates Piezo1 channels in brain capillary endothelial cells. . [ Google Scholar ] 320. Thakore P, Yamasaki E, Ali S, Sanchez Solano A, Labelle-Dumais C, Gao X, Chaumeil MM, Gould DB, Earley S. PI3K block restores age-dependent neurovascular coupling defects associated with cerebral small vessel disease. Proc Natl Acad Sci U S A 120: e2306479120, 2023. doi: 10.1073/pnas.2306479120. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 321. The Axon Guide, Electrophysiology and Biophysics Laboratory Techniques. Molecular Devices, 2012. [ Google Scholar ] 322. Huang S, Uusisaari MY. Physiological temperature during brain slicing enhances the quality of acute slice preparations. Front Cell Neurosci 7: 48, 2013. doi: 10.3389/fncel.2013.00048. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 323. Longden TA, Dabertrand F, Koide M, Gonzales AL, Tykocki NR, Brayden JE, Hill-Eubanks D, Nelson MT. Capillary K+-sensing initiates retrograde hyperpolarization to increase local cerebral blood flow. Nat Neurosci 20: 717–726, 2017. doi: 10.1038/nn.4533. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 324. Tajada S, Moreno CM, O’Dwyer S, Woods S, Sato D, Navedo MF, Santana LF. Distance constraints on activation of TRPV4 channels by AKAP150-bound PKCα in arterial myocytes. J Gen Physiol 149: 639–659, 2017. doi: 10.1085/jgp.201611709. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 325. Rae J, Cooper K, Gates P, Watsky M. Low access resistance perforated patch recordings using amphotericin B. J Neurosci Methods 37: 15–26, 1991. doi: 10.1016/0165-0270(91)90017-t. [ DOI ] [ PubMed ] [ Google Scholar ] 326. Horn R, Marty A. Muscarinic activation of ionic currents measured by a new whole-cell recording method. J Gen Physiol 92: 145–59, 1988. doi: 10.1085/jgp.92.2.145. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 327. Gonzales AL, Yang Y, Sullivan MN, Sanders L, Dabertrand F, Hill-Eubanks DC, Nelson MT, Earley S. A PLCγ1-dependent, force-sensitive signaling network in the myogenic constriction of cerebral arteries. Sci Signal 7: ra49, 2014. doi: 10.1126/scisignal.2004732. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 328. Abe Y, Furukawa K, Itoyama Y, Akaike N. Glycine response in acutely dissociated ventromedial hypothalamic neuron of the rat: new approach with gramicidin perforated patch-clamp technique. J Neurophysiol 72: 1530–7, 1994. doi: 10.1152/jn.1994.72.4.1530. [ DOI ] [ PubMed ] [ Google Scholar ] 329. von Beckerath N, Dittrich M, Klieber HG, Daut J. Inwardly rectifying K+ channels in freshly dissociated coronary endothelial cells from guinea-pig heart. J Physiol 491 (Pt 2): 357–65, 1996. doi: 10.1113/jphysiol.1996.sp021221. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 330. Yamasaki E, Ali S, Sanchez Solano A, Thakore P, Smith M, Wang X, Labelle-Dumais C, Gould DB, Earley S. Faulty TRPM4 channels underlie age-dependent cerebral vascular dysfunction in Gould syndrome. Proc Natl Acad Sci U S A 120: e2217327120, 2023. doi: 10.1073/pnas.2217327120. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 331. Mata-Daboin A, Garrud TAC, Fernandez-Pena C, Peixoto-Neves D, Leo MD, Bernardelli AK, Singh P, Malik KU, Jaggar JH. Vasodilators activate the anion channel TMEM16A in endothelial cells to reduce blood pressure. Sci Signal 16: eadh9399, 2023. doi: 10.1126/scisignal.adh9399. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 332. Lenaeus MJ, Vamvouka M, Focia PJ, Gross A. Structural basis of TEA blockade in a model potassium channel. Nat Struct Mol Biol 12: 454–9, 2005. doi: 10.1038/nsmb929. [ DOI ] [ PubMed ] [ Google Scholar ] 333. Sonkusare SK, Dalsgaard T, Bonev AD, Nelson MT. Inward rectifier potassium (Kir2.1) channels as end-stage boosters of endothelium-dependent vasodilators. J Physiol 594: 3271–85, 2016. doi: 10.1113/JP271652. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 334. Spahiu E, Kastrati E, Amrute-Nayak M. PyChelator: a Python-based Colab and web application for metal chelator calculations. BMC Bioinformatics 25: 239, 2024. doi: 10.1186/s12859-024-05858-8. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 335. He W-Q, Qiao Y-N, Zhang C-H, Peng Y-J, Chen C, Wang P, Gao Y-Q, Chen C, Chen X, Tao T, Su X-H, Li C-J, Kamm KE, Stull JT, Zhu M-S. Role of myosin light chain kinase in regulation of basal blood pressure and maintenance of salt-induced hypertension. Am J Physiol Heart Circ Physiol 301: H584–91, 2011. doi: 10.1152/ajpheart.01212.2010. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 336. Thakore P, Alvarado MG, Ali S, Mughal A, Pires PW, Yamasaki E, Pritchard HA, Isakson BE, Tran CHT, Earley S. Brain endothelial cell TRPA1 channels initiate neurovascular coupling. Elife 10, 2021. doi: 10.7554/eLife.63040. [ DOI ] [ Google Scholar ] 337. Ledoux J, Bonev AD, Nelson MT. Ca2+-activated K+ channels in murine endothelial cells: block by intracellular calcium and magnesium. J Gen Physiol 131: 125–35, 2008. doi: 10.1085/jgp.200709875. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 338. Bagher P, Polo-Parada L, Segal SS. Microiontophoresis and micromanipulation for intravital fluorescence imaging of the microcirculation. J Vis Exp , 2011. doi: 10.3791/2900. [ DOI ] [ Google Scholar ] 339. Marino M, Misuri L, Brogioli D. A new open source software for the calculation of the liquid junction potential between two solutions according to the stationary Nernst-Planck equation. [ Google Scholar ] 340. Nilius B, Viana F, Droogmans G. Ion channels in vascular endothelium. Annu Rev Physiol 59: 145–70, 1997. doi: 10.1146/annurev.physiol.59.1.145. [ DOI ] [ PubMed ] [ Google Scholar ] 341. Barbee KA, Davies PF, Lal R. Shear stress-induced reorganization of the surface topography of living endothelial cells imaged by atomic force microscopy. Circ Res 74: 163–71, 1994. doi: 10.1161/01.res.74.1.163. [ DOI ] [ PubMed ] [ Google Scholar ] 342. Byfield FJ, Reen RK, Shentu T-P, Levitan I, Gooch KJ. Endothelial actin and cell stiffness is modulated by substrate stiffness in 2D and 3D. J Biomech 42: 1114–9, 2009. doi: 10.1016/j.jbiomech.2009.02.012. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 343. Wiesinger A, Peters W, Chappell D, Kentrup D, Reuter S, Pavenstädt H, Oberleithner H, Kümpers P. Nanomechanics of the endothelial glycocalyx in experimental sepsis. PLoS One 8: e80905, 2013. doi: 10.1371/journal.pone.0080905. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 344. Targosz-Korecka M, Jaglarz M, Malek-Zietek KE, Gregorius A, Zakrzewska A, Sitek B, Rajfur Z, Chlopicki S, Szymonski M. AFM-based detection of glycocalyx degradation and endothelial stiffening in the db/db mouse model of diabetes. Sci Rep 7: 15951, 2017. doi: 10.1038/s41598-017-16179-7. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 345. Martinez-Lemus LA, Sun Z, Trache A, Trzciakowski JP, Meininger GA. Integrins and regulation of the microcirculation: from arterioles to molecular studies using atomic force microscopy. Microcirculation 12: 99–112, 2005. doi: 10.1080/10739680590896054. [ DOI ] [ PubMed ] [ Google Scholar ] 346. DeMarco VG, Habibi J, Jia G, Aroor AR, Ramirez-Perez FI, Martinez-Lemus LA, Bender SB, Garro M, Hayden MR, Sun Z, Meininger GA, Manrique C, Whaley-Connell A, Sowers JR. Low-Dose Mineralocorticoid Receptor Blockade Prevents Western Diet-Induced Arterial Stiffening in Female Mice. Hypertension 66: 99–107, 2015. doi: 10.1161/HYPERTENSIONAHA.115.05674. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 347. McCallinhart PE, Cho Y, Sun Z, Ghadiali S, Meininger GA, Trask AJ. Reduced stiffness and augmented traction force in type 2 diabetic coronary microvascular smooth muscle. Am J Physiol Heart Circ Physiol 318: H1410–H1419, 2020. doi: 10.1152/ajpheart.00542.2019. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 348. Efremov YM, Dokrunova AA, Bagrov DV, Kudryashova KS, Sokolova OS, Shaitan KV. The effects of confluency on cell mechanical properties. J Biomech 46: 1081–7, 2013. doi: 10.1016/j.jbiomech.2013.01.022. [ DOI ] [ PubMed ] [ Google Scholar ] 349. Gavara N. A beginner’s guide to atomic force microscopy probing for cell mechanics. Microsc Res Tech 80: 75–84, 2017. doi: 10.1002/jemt.22776. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 350. Caille N, Thoumine O, Tardy Y, Meister J-J. Contribution of the nucleus to the mechanical properties of endothelial cells. J Biomech 35: 177–87, 2002. doi: 10.1016/s0021-9290(01)00201-9. [ DOI ] [ PubMed ] [ Google Scholar ] 351. Akhtar R, Schwarzer N, Sherratt MJ, Watson REB, Graham HK, Trafford AW, Mummery PM, Derby B. Nanoindentation of histological specimens: Mapping the elastic properties of soft tissues. J Mater Res 24: 638–646, 2009. doi: 10.1557/JMR.2009.0130. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 352. Han Y, Cho Y-E, Ayon R, Guo R, Youssef KD, Pan M, Dai A, Yuan JX- J, Makino A. SGLT inhibitors attenuate NO-dependent vascular relaxation in the pulmonary artery but not in the coronary artery. Am J Physiol Lung Cell Mol Physiol 309: L1027–36, 2015. doi: 10.1152/ajplung.00167.2015. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 353. Tumova S, Kerimi A, Porter KE, Williamson G. Transendothelial glucose transport is not restricted by extracellular hyperglycaemia. Vascul Pharmacol 87: 219–229, 2016. doi: 10.1016/j.vph.2016.11.001. [ DOI ] [ PubMed ] [ Google Scholar ] 354. Dobrina A, Rossi F. Metabolic properties of freshly isolated bovine endothelial cells. Biochim Biophys Acta 762: 295–301, 1983. doi: 10.1016/0167-4889(83)90084-8. [ DOI ] [ PubMed ] [ Google Scholar ] 355. Krützfeldt A, Spahr R, Mertens S, Siegmund B, Piper HM. Metabolism of exogenous substrates by coronary endothelial cells in culture. J Mol Cell Cardiol 22: 1393–404, 1990. doi: 10.1016/0022-2828(90)90984-a. [ DOI ] [ PubMed ] [ Google Scholar ] 356. De Bock K, Georgiadou M, Schoors S, Kuchnio A, Wong BW, Cantelmo AR, Quaegebeur A, Ghesquière B, Cauwenberghs S, Eelen G, Phng L-K, Betz I, Tembuyser B, Brepoels K, Welti J, Geudens I, Segura I, Cruys B, Bifari F, Decimo I, Blanco R, Wyns S, Vangindertael J, Rocha S, Collins RT, Munck S, Daelemans D, Imamura H, Devlieger R, Rider M, Van Veldhoven PP, Schuit F, Bartrons R, Hofkens J, Fraisl P, Telang S, Deberardinis RJ, Schoonjans L, Vinckier S, Chesney J, Gerhardt H, Dewerchin M, Carmeliet P. Role of PFKFB3-driven glycolysis in vessel sprouting. Cell 154: 651–63, 2013. doi: 10.1016/j.cell.2013.06.037. [ DOI ] [ PubMed ] [ Google Scholar ] 357. Rigoulet M, Bouchez CL, Paumard P, Ransac S, Cuvellier S, Duvezin-Caubet S, Mazat JP, Devin A. Cell energy metabolism: An update. Biochim Biophys Acta Bioenerg 1861: 148276, 2020. doi: 10.1016/j.bbabio.2020.148276. [ DOI ] [ PubMed ] [ Google Scholar ] 358. Groschner LN, Waldeck-Weiermair M, Malli R, Graier WF. Endothelial mitochondria--less respiration, more integration. Pflugers Arch 464: 63–76, 2012. doi: 10.1007/s00424-012-1085-z. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 359. Yang M, Chadwick AE, Dart C, Kamishima T, Quayle JM. Bioenergetic profile of human coronary artery smooth muscle cells and effect of metabolic intervention. PLoS One 12: e0177951, 2017. doi: 10.1371/journal.pone.0177951. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 360. Leite-Moreira AF, Rocha-Sousa A, Henriques-Coelho T. Cardiac, skeletal, and smooth muscle regulation by ghrelin. Vitam Horm 77: 207–38, 2008. doi: 10.1016/S0083-6729(06)77009-1. [ DOI ] [ PubMed ] [ Google Scholar ] 361. Panda K, Adak S, Konas D, Sharma M, Stuehr DJ. A conserved aspartate (Asp-1393) regulates NADPH reduction of neuronal nitric-oxide synthase: implications for catalysis. J Biol Chem 279: 18323–33, 2004. doi: 10.1074/jbc.M310391200. [ DOI ] [ PubMed ] [ Google Scholar ] 362. Kruger NJ, von Schaewen A. The oxidative pentose phosphate pathway: structure and organisation. Curr Opin Plant Biol 6: 236–46, 2003. doi: 10.1016/s1369-5266(03)00039-6. [ DOI ] [ PubMed ] [ Google Scholar ] 363. Haltiwanger RS, Kelly WG, Roquemore EP, Blomberg MA, Dong LY, Kreppel L, Chou TY, Hart GW. Glycosylation of nuclear and cytoplasmic proteins is ubiquitous and dynamic. Biochem Soc Trans 20: 264–9, 1992. doi: 10.1042/bst0200264. [ DOI ] [ PubMed ] [ Google Scholar ] 364. Horton RW, Meldrum BS, Bachelard HS. Enzymic and cerebral metabolic effects of 2-deoxy-D-glucose. J Neurochem 21: 507–20, 1973. doi: 10.1111/j.1471-4159.1973.tb05996.x. [ DOI ] [ PubMed ] [ Google Scholar ] 365. Tao J, Diaz RK, Teixeira CR V, Hackmann TJ. Transport of a Fluorescent Analogue of Glucose (2-NBDG) versus Radiolabeled Sugars by Rumen Bacteria and Escherichia coli. Biochemistry 55: 2578–89, 2016. doi: 10.1021/acs.biochem.5b01286. [ DOI ] [ PubMed ] [ Google Scholar ] 366. Zheng Q, Cabrera JTO, Tsuji-Hosokawa A, Ramirez FJ, Cai H, Yuan JX- J, Wang J, Makino A. Enhanced lung endothelial glycolysis is implicated in the development of severe pulmonary hypertension in type 2 diabetes. Am J Physiol Lung Cell Mol Physiol 328: L430–L442, 2025. doi: 10.1152/ajplung.00305.2023. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 367. Schönfeld P, Wojtczak L. Short- and medium-chain fatty acids in energy metabolism: the cellular perspective. J Lipid Res 57: 943–54, 2016. doi: 10.1194/jlr.R067629. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 368. Bailey JM, Howard BV, Tillman SF. Lipid metabolism in cultured cells. XI. Utilization of serum triglycerides. J Biol Chem 248: 1240–7, 1973. [ PubMed ] [ Google Scholar ] 369. Chandel NS. Amino Acid Metabolism. Cold Spring Harb Perspect Biol 13, 2021. doi: 10.1101/cshperspect.a040584. [ DOI ] [ Google Scholar ] 370. Hewton KG, Johal AS, Parker SJ. Transporters at the Interface between Cytosolic and Mitochondrial Amino Acid Metabolism. Metabolites 11, 2021. doi: 10.3390/metabo11020112. [ DOI ] [ Google Scholar ] 371. Lorenz M, Fritsche-Guenther R, Bartsch C, Vietzke A, Eisenberger A, Stangl K, Stangl V, Kirwan JA. Serum Starvation Accelerates Intracellular Metabolism in Endothelial Cells. Int J Mol Sci 24, 2023. doi: 10.3390/ijms24021189. [ DOI ] [ Google Scholar ] 372. Zhang W, Li H, Ogando DG, Li S, Feng M, Price FW, Tennessen JM, Bonanno JA. Glutaminolysis is Essential for Energy Production and Ion Transport in Human Corneal Endothelium. EBioMedicine 16: 292–301, 2017. doi: 10.1016/j.ebiom.2017.01.004. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 373. Schoors S, Bruning U, Missiaen R, Queiroz KC, Borgers G, Elia I, Zecchin A, Cantelmo AR, Christen S, Goveia J, Heggermont W, Goddé L, Vinckier S, Van Veldhoven PP, Eelen G, Schoonjans L, Gerhardt H, Dewerchin M, Baes M, De Bock K, Ghesquière B, Lunt SY, Fendt S-M, Carmeliet P. Fatty acid carbon is essential for dNTP synthesis in endothelial cells. Nature 520: 192–197, 2015. doi: 10.1038/nature14362. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 374. Alseekh S, Aharoni A, Brotman Y, Contrepois K, D’Auria J, Ewald J, C Ewald J, Fraser PD, Giavalisco P, Hall RD, Heinemann M, Link H, Luo J, Neumann S, Nielsen J, Perez de Souza L, Saito K, Sauer U, Schroeder FC, Schuster S, Siuzdak G, Skirycz A, Sumner LW, Snyder MP, Tang H, Tohge T, Wang Y, Wen W, Wu S, Xu G, Zamboni N, Fernie AR. Mass spectrometry-based metabolomics: a guide for annotation, quantification and best reporting practices. Nat Methods 18: 747–756, 2021. doi: 10.1038/s41592-021-01197-1. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 375. Nagana Gowda GA, Raftery D. NMR-Based Metabolomics. Adv Exp Med Biol 1280: 19–37, 2021. doi: 10.1007/978-3-030-51652-9_2. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 376. Förstermann U, Sessa WC. Nitric oxide synthases: regulation and function. Eur Heart J 33: 829-37, 837a–837d, 2012. doi: 10.1093/eurheartj/ehr304. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 377. Münzel T, Daiber A. Vascular Redox Signaling, Endothelial Nitric Oxide Synthase Uncoupling, and Endothelial Dysfunction in the Setting of Transportation Noise Exposure or Chronic Treatment with Organic Nitrates. Antioxid Redox Signal 38: 1001–1021, 2023. doi: 10.1089/ars.2023.0006. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 378. Zhu L, Zhang Y, Guo Z, Wang M. Cardiovascular Biology of Prostanoids and Drug Discovery. Arterioscler Thromb Vasc Biol 40: 1454–1463, 2020. doi: 10.1161/ATVBAHA.119.313234. [ DOI ] [ PubMed ] [ Google Scholar ] 379. Vanhoutte PM, Shimokawa H, Feletou M, Tang EHC. Endothelial dysfunction and vascular disease - a 30th anniversary update. Acta Physiol (Oxf) 219: 22–96, 2017. doi: 10.1111/apha.12646. [ DOI ] [ PubMed ] [ Google Scholar ] 380. Hernanz R, Briones AM, Salaices M, Alonso MJ. New roles for old pathways? A circuitous relationship between reactive oxygen species and cyclo-oxygenase in hypertension. Clin Sci (Lond) 126: 111–21, 2014. doi: 10.1042/CS20120651. [ DOI ] [ PubMed ] [ Google Scholar ] 381. Majed BH, Khalil RA. Molecular mechanisms regulating the vascular prostacyclin pathways and their adaptation during pregnancy and in the newborn. Pharmacol Rev 64: 540–82, 2012. doi: 10.1124/pr.111.004770. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 382. Jones RL, Giembycz MA, Woodward DF. Prostanoid receptor antagonists: development strategies and therapeutic applications. Br J Pharmacol 158: 104–45, 2009. doi: 10.1111/j.1476-5381.2009.00317.x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 383. Mazaleuskaya LL, Ricciotti E. Druggable Prostanoid Pathway. Adv Exp Med Biol 1274: 29–54, 2020. doi: 10.1007/978-3-030-50621-6_3. [ DOI ] [ PubMed ] [ Google Scholar ] 384. Tang EHC, Vanhoutte PM. Prostanoids and reactive oxygen species: team players in endothelium-dependent contractions. Pharmacol Ther 122: 140–9, 2009. doi: 10.1016/j.pharmthera.2009.02.006. [ DOI ] [ PubMed ] [ Google Scholar ] 385. Camargo LL, Rios FJ, Montezano AC, Touyz RM. Reactive oxygen species in hypertension. Nat Rev Cardiol 22: 20–37, 2025. doi: 10.1038/s41569-024-01062-6. [ DOI ] [ PubMed ] [ Google Scholar ] 386. Casas AI, Nogales C, Mucke HAM, Petraina A, Cuadrado A, Rojo AI, Ghezzi P, Jaquet V, Augsburger F, Dufrasne F, Soubhye J, Deshwal S, Di Sante M, Kaludercic N, Di Lisa F, Schmidt HHHW. On the Clinical Pharmacology of Reactive Oxygen Species. Pharmacol Rev 72: 801–828, 2020. doi: 10.1124/pr.120.019422. [ DOI ] [ PubMed ] [ Google Scholar ] 387. Forman HJ, Zhang H. Targeting oxidative stress in disease: promise and limitations of antioxidant therapy. Nat Rev Drug Discov 20: 689–709, 2021. doi: 10.1038/s41573-021-00233-1. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 388. Pagano PJ, Cifuentes-Pagano E. The Enigmatic Vascular NOX: From Artifact to Double Agent of Change: Arthur C. Corcoran Memorial Lecture - 2019. Hypertension 77: 275–283, 2021. doi: 10.1161/HYPERTENSIONAHA.120.13897. [ DOI ] [ PubMed ] [ Google Scholar ] 389. Dao VT- V, Elbatreek MH, Altenhöfer S, Casas AI, Pachado MP, Neullens CT, Knaus UG, Schmidt HHHW. Isoform-selective NADPH oxidase inhibitor panel for pharmacological target validation. Free Radic Biol Med 148: 60–69, 2020. doi: 10.1016/j.freeradbiomed.2019.12.038. [ DOI ] [ PubMed ] [ Google Scholar ] 390. Cipriano A, Viviano M, Feoli A, Milite C, Sarno G, Castellano S, Sbardella G. NADPH Oxidases: From Molecular Mechanisms to Current Inhibitors. J Med Chem 66: 11632–11655, 2023. doi: 10.1021/acs.jmedchem.3c00770. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 391. Cuadrado A, Cazalla E, Bach A, Bathish B, Naidu SD, DeNicola GM, Dinkova-Kostova AT, Fernández-Ginés R, Grochot-Przeczek A, Hayes JD, Kensler TW, León R, Liby KT, López MG, Manda G, Shivakumar AK, Hakomäki H, Moerland JA, Motohashi H, Rojo AI, Sykiotis GP, Taguchi K, Valverde ÁM, Yamamoto M, Levonen A-L. Health position paper and redox perspectives - Bench to bedside transition for pharmacological regulation of NRF2 in noncommunicable diseases. Redox Biol 81: 103569, 2025. doi: 10.1016/j.redox.2025.103569. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 392. Liu T, Zhang M, Mukosera GT, Borchardt D, Li Q, Tipple TE, Ishtiaq Ahmed AS, Power GG, Blood AB. L-NAME releases nitric oxide and potentiates subsequent nitroglycerin-mediated vasodilation. Redox Biol 26: 101238, 2019. doi: 10.1016/j.redox.2019.101238. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 393. Garvey EP, Oplinger JA, Furfine ES, Kiff RJ, Laszlo F, Whittle BJ, Knowles RG. 1400W is a slow, tight binding, and highly selective inhibitor of inducible nitric-oxide synthase in vitro and in vivo. J Biol Chem 272: 4959–63, 1997. doi: 10.1074/jbc.272.8.4959. [ DOI ] [ PubMed ] [ Google Scholar ] 394. Szabó C, Southan GJ, Thiemermann C. Beneficial effects and improved survival in rodent models of septic shock with S-methylisothiourea sulfate, a potent and selective inhibitor of inducible nitric oxide synthase. Proc Natl Acad Sci U S A 91: 12472–6, 1994. doi: 10.1073/pnas.91.26.12472. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 395. Smith CJ, Zhang Y, Koboldt CM, Muhammad J, Zweifel BS, Shaffer A, Talley JJ, Masferrer JL, Seibert K, Isakson PC. Pharmacological analysis of cyclooxygenase-1 in inflammation. Proc Natl Acad Sci U S A 95: 13313–8, 1998. doi: 10.1073/pnas.95.22.13313. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 396. Brenneis C, Maier TJ, Schmidt R, Hofacker A, Zulauf L, Jakobsson P-J, Scholich K, Geisslinger G. Inhibition of prostaglandin E2 synthesis by SC-560 is independent of cyclooxygenase 1 inhibition. FASEB J 20: 1352–60, 2006. doi: 10.1096/fj.05-5346com. [ DOI ] [ PubMed ] [ Google Scholar ] 397. Futaki N, Takahashi S, Yokoyama M, Arai I, Higuchi S, Otomo S. NS-398, a new anti-inflammatory agent, selectively inhibits prostaglandin G/H synthase/cyclooxygenase (COX-2) activity in vitro. Prostaglandins 47: 55–9, 1994. doi: 10.1016/0090-6980(94)90074-4. [ DOI ] [ PubMed ] [ Google Scholar ] 398. Riendeau D, Percival MD, Boyce S, Brideau C, Charleson S, Cromlish W, Ethier D, Evans J, Falgueyret JP, Ford-Hutchinson AW, Gordon R, Greig G, Gresser M, Guay J, Kargman S, Léger S, Mancini JA, O’Neill G, Ouellet M, Rodger IW, Thérien M, Wang Z, Webb JK, Wong E, Chan CC. Biochemical and pharmacological profile of a tetrasubstituted furanone as a highly selective COX-2 inhibitor. Br J Pharmacol 121: 105–17, 1997. doi: 10.1038/sj.bjp.0701076. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 399. Leclerc P, Idborg H, Spahiu L, Larsson C, Nekhotiaeva N, Wannberg J, Stenberg P, Korotkova M, Jakobsson P-J. Characterization of a human and murine mPGES-1 inhibitor and comparison to mPGES-1 genetic deletion in mouse models of inflammation. Prostaglandins Other Lipid Mediat 107: 26–34, 2013. doi: 10.1016/j.prostaglandins.2013.09.001. [ DOI ] [ PubMed ] [ Google Scholar ] 400. Larsson K, Steinmetz J, Bergqvist F, Arefin S, Spahiu L, Wannberg J, Pawelzik S-C, Morgenstern R, Stenberg P, Kublickiene K, Korotkova M, Jakobsson P-J. Biological characterization of new inhibitors of microsomal PGE synthase-1 in preclinical models of inflammation and vascular tone. Br J Pharmacol 176: 4625–4638, 2019. doi: 10.1111/bph.14827. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 401. Bley KR, Bhattacharya A, Daniels DV, Gever J, Jahangir A, O’Yang C, Smith S, Srinivasan D, Ford APDW, Jett M-F. RO1138452 and RO3244794: characterization of structurally distinct, potent and selective IP (prostacyclin) receptor antagonists. Br J Pharmacol 147: 335–45, 2006. doi: 10.1038/sj.bjp.0706554. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 402. af Forselles KJ, Root J, Clarke T, Davey D, Aughton K, Dack K, Pullen N. In vitro and in vivo characterization of PF-04418948, a novel, potent and selective prostaglandin EP 2 receptor antagonist. Br J Pharmacol 164: 1847–56, 2011. doi: 10.1111/j.1476-5381.2011.01495.x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 403. Wilcox CS. Effects of tempol and redox-cycling nitroxides in models of oxidative stress. Pharmacol Ther 126: 119–45, 2010. doi: 10.1016/j.pharmthera.2010.01.003. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 404. Simonsen U, Christensen FH, Buus NH. The effect of tempol on endothelium-dependent vasodilatation and blood pressure. Pharmacol Ther 122: 109–24, 2009. doi: 10.1016/j.pharmthera.2009.02.002. [ DOI ] [ PubMed ] [ Google Scholar ] 405. Pedre B, Barayeu U, Ezeriņa D, Dick TP. The mechanism of action of N-acetylcysteine (NAC): The emerging role of H2S and sulfane sulfur species. Pharmacol Ther 228: 107916, 2021. doi: 10.1016/j.pharmthera.2021.107916. [ DOI ] [ PubMed ] [ Google Scholar ] 406. Sarvani C, Sireesh D, Ramkumar KM. Unraveling the role of ER stress inhibitors in the context of metabolic diseases. Pharmacol Res 119: 412–421, 2017. doi: 10.1016/j.phrs.2017.02.018. [ DOI ] [ PubMed ] [ Google Scholar ] 407. Heumüller S, Wind S, Barbosa-Sicard E, Schmidt HHHW, Busse R, Schröder K, Brandes RP. Apocynin is not an inhibitor of vascular NADPH oxidases but an antioxidant. Hypertension 51: 211–7, 2008. doi: 10.1161/HYPERTENSIONAHA.107.100214. [ DOI ] [ PubMed ] [ Google Scholar ] 408. Augsburger F, Filippova A, Rasti D, Seredenina T, Lam M, Maghzal G, Mahiout Z, Jansen-Dürr P, Knaus UG, Doroshow J, Stocker R, Krause K-H, Jaquet V. Pharmacological characterization of the seven human NOX isoforms and their inhibitors. Redox Biol 26: 101272, 2019. doi: 10.1016/j.redox.2019.101272. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 409. Ranayhossaini DJ, Rodriguez AI, Sahoo S, Chen BB, Mallampalli RK, Kelley EE, Csanyi G, Gladwin MT, Romero G, Pagano PJ. Selective recapitulation of conserved and nonconserved regions of putative NOXA1 protein activation domain confers isoform-specific inhibition of Nox1 oxidase and attenuation of endothelial cell migration. J Biol Chem 288: 36437–50, 2013. doi: 10.1074/jbc.M113.521344. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 410. Csányi G, Cifuentes-Pagano E, Al Ghouleh I, Ranayhossaini DJ, Egaña L, Lopes LR, Jackson HM, Kelley EE, Pagano PJ. Nox2 B-loop peptide, Nox2ds, specifically inhibits the NADPH oxidase Nox2. Free Radic Biol Med 51: 1116–25, 2011. doi: 10.1016/j.freeradbiomed.2011.04.025. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 411. Luo J, Yan D, Li S, Liu S, Zeng F, Cheung CW, Liu H, Irwin MG, Huang H, Xia Z. Allopurinol reduces oxidative stress and activates Nrf2/p62 to attenuate diabetic cardiomyopathy in rats. J Cell Mol Med 24: 1760–1773, 2020. doi: 10.1111/jcmm.14870. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 412. Dikalova AE, Bikineyeva AT, Budzyn K, Nazarewicz RR, McCann L, Lewis W, Harrison DG, Dikalov SI. Therapeutic targeting of mitochondrial superoxide in hypertension. Circ Res 107: 106–16, 2010. doi: 10.1161/CIRCRESAHA.109.214601. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 413. Escribano-Lopez I, Diaz-Morales N, Rovira-Llopis S, de Marañon AM, Orden S, Alvarez A, Bañuls C, Rocha M, Murphy MP, Hernandez-Mijares A, Victor VM. The mitochondria-targeted antioxidant MitoQ modulates oxidative stress, inflammation and leukocyte-endothelium interactions in leukocytes isolated from type 2 diabetic patients. Redox Biol 10: 200–205, 2016. doi: 10.1016/j.redox.2016.10.017. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 414. Battson ML, Lee DM, Gentile CL. Endoplasmic reticulum stress and the development of endothelial dysfunction. Am J Physiol Heart Circ Physiol 312: H355–H367, 2017. doi: 10.1152/ajpheart.00437.2016. [ DOI ] [ PubMed ] [ Google Scholar ] 415. Dinkova-Kostova AT, Copple IM. Advances and challenges in therapeutic targeting of NRF2. Trends Pharmacol Sci 44: 137–149, 2023. doi: 10.1016/j.tips.2022.12.003. [ DOI ] [ PubMed ] [ Google Scholar ] 416. Ellinsworth DC, Sandow SL, Shukla N, Liu Y, Jeremy JY, Gutterman DD. Endothelium-Derived Hyperpolarization and Coronary Vasodilation: Diverse and Integrated Roles of Epoxyeicosatrienoic Acids, Hydrogen Peroxide, and Gap Junctions. Microcirculation 23: 15–32, 2016. doi: 10.1111/micc.12255. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 417. Garland CJ, Dora KA. EDH: endothelium-dependent hyperpolarization and microvascular signalling. Acta Physiol (Oxf) 219: 152–161, 2017. doi: 10.1111/apha.12649. [ DOI ] [ PubMed ] [ Google Scholar ] 418. Garland CJ, McPherson GA. Evidence that nitric oxide does not mediate the hyperpolarization and relaxation to acetylcholine in the rat small mesenteric artery. Br J Pharmacol 105: 429–35, 1992. doi: 10.1111/j.1476-5381.1992.tb14270.x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 419. Crane GJ, Gallagher N, Dora KA, Garland CJ. Small- and intermediate-conductance calcium-activated K+ channels provide different facets of endothelium-dependent hyperpolarization in rat mesenteric artery. J Physiol 553: 183–9, 2003. doi: 10.1113/jphysiol.2003.051896. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 420. Edwards G, Dora KA, Gardener MJ, Garland CJ, Weston AH. K+ is an endothelium-derived hyperpolarizing factor in rat arteries. Nature 396: 269–72, 1998. doi: 10.1038/24388. [ DOI ] [ PubMed ] [ Google Scholar ] 421. Boedtkjer E, Kim S, Aalkjaer C. Endothelial alkalinisation inhibits gap junction communication and endothelium-derived hyperpolarisations in mouse mesenteric arteries. J Physiol 591: 1447–61, 2013. doi: 10.1113/jphysiol.2012.247478. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 422. Segal SS, Bény JL. Intracellular recording and dye transfer in arterioles during blood flow control. Am J Physiol 263: H1–7, 1992. doi: 10.1152/ajpheart.1992.263.1.H1. [ DOI ] [ PubMed ] [ Google Scholar ] 423. Dora KA, Garland CJ. Properties of smooth muscle hyperpolarization and relaxation to K+ in the rat isolated mesenteric artery. Am J Physiol Heart Circ Physiol 280: H2424–9, 2001. doi: 10.1152/ajpheart.2001.280.6.H2424. [ DOI ] [ PubMed ] [ Google Scholar ] 424. Dora KA, Ings NT, Garland CJ. K(Ca) channel blockers reveal hyperpolarization and relaxation to K+ in rat isolated mesenteric artery. Am J Physiol Heart Circ Physiol 283: H606–14, 2002. doi: 10.1152/ajpheart.01016.2001. [ DOI ] [ PubMed ] [ Google Scholar ] 425. Leurgans TM, Bloksgaard M, Brewer JR, Bagatolli LA, Fredgart MH, Rosenstand K, Hansen ML, Rasmussen LM, Irmukhamedov A, De Mey JG. Endothelin-1 shifts the mediator of bradykinin-induced relaxation from NO to H2 O2 in resistance arteries from patients with cardiovascular disease. Br J Pharmacol 173: 1653–64, 2016. doi: 10.1111/bph.13467. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 426. Leurgans TM, Bloksgaard M, Irmukhamedov A, Riber LP, De Mey JGR. Relaxing Responses to Hydrogen Peroxide and Nitric Oxide in Human Pericardial Resistance Arteries Stimulated with Endothelin-1. Basic Clin Pharmacol Toxicol 122: 74–81, 2018. doi: 10.1111/bcpt.12843. [ DOI ] [ PubMed ] [ Google Scholar ] 427. Crane GJ, Garland CJ. Thromboxane receptor stimulation associated with loss of SKCa activity and reduced EDHF responses in the rat isolated mesenteric artery. Br J Pharmacol 142: 43–50, 2004. doi: 10.1038/sj.bjp.0705756. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 428. Brown BM, Shim H, Christophersen P, Wulff H. Pharmacology of Small- and Intermediate-Conductance Calcium-Activated Potassium Channels. Annu Rev Pharmacol Toxicol 60: 219–240, 2020. doi: 10.1146/annurev-pharmtox-010919-023420. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 429. Mombouli JV, Vanhoutte PM. Heterogeneity of endothelium-dependent vasodilator effects of angiotensin-converting enzyme inhibitors: role of bradykinin generation during ACE inhibition. J Cardiovasc Pharmacol 20 Suppl 9: S74–82, 1992. [ Google Scholar ] 430. Hansen FB, Esteves GV, Mogensen S, Prat-Duran J, Secher N, Løfgren B, Granfeldt A, Simonsen U. Increased cerebral endothelium-dependent vasodilation in rats in the postcardiac arrest period. J Appl Physiol (1985) 131: 1311–1327, 2021. doi: 10.1152/japplphysiol.00373.2021. [ DOI ] [ PubMed ] [ Google Scholar ] 431. Pomposiello S, Rhaleb NE, Alva M, Carretero OA. Reactive oxygen species: role in the relaxation induced by bradykinin or arachidonic acid via EDHF in isolated porcine coronary arteries. J Cardiovasc Pharmacol 34: 567–74, 1999. doi: 10.1097/00005344-199910000-00014. [ DOI ] [ PubMed ] [ Google Scholar ] 432. White TD, Angus JA. Relaxant effects of ATP and adenosine on canine large and small coronary arteries in vitro. Eur J Pharmacol 143: 119–26, 1987. doi: 10.1016/0014-2999(87)90741-2. [ DOI ] [ PubMed ] [ Google Scholar ] 433. King AD, Milavec-Krizman M, Müller-Schweinitzer E. Characterization of the adenosine receptor in porcine coronary arteries. Br J Pharmacol 100: 483–6, 1990. doi: 10.1111/j.1476-5381.1990.tb15833.x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 434. Meens MJPMT, Compeer MG, Hackeng TM, van Zandvoort MA, Janssen BJA, De Mey JGR. Stimuli of sensory-motor nerves terminate arterial contractile effects of endothelin-1 by CGRP and dissociation of ET-1/ET(A)-receptor complexes. PLoS One 5: e10917, 2010. doi: 10.1371/journal.pone.0010917. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 435. Durand MJ, Gutterman DD. Diversity in mechanisms of endothelium-dependent vasodilation in health and disease. Microcirculation 20: 239–47, 2013. doi: 10.1111/micc.12040. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 436. Wang X, He B. Endothelial dysfunction: molecular mechanisms and clinical implications. MedComm (Beijing) 5: e651, 2024. doi: 10.1002/mco2.651. [ DOI ] [ Google Scholar ] 437. Jaiswal N, Lambrecht G, Mutschler E, Tacke R, Malik KU. Pharmacological characterization of the vascular muscarinic receptors mediating relaxation and contraction in rabbit aorta. J Pharmacol Exp Ther 258: 842–50, 1991. [ PubMed ] [ Google Scholar ] 438. Eglen RM, Reddy H, Watson N, Challiss RA. Muscarinic acetylcholine receptor subtypes in smooth muscle. Trends Pharmacol Sci 15: 114–9, 1994. doi: 10.1016/0165-6147(94)90047-7. [ DOI ] [ PubMed ] [ Google Scholar ] 439. Patil PN, Stearns R. Mechanism of vascular relaxation by cholinomimetic drugs with special reference to pilocarpine and arecoline. J Ocul Pharmacol Ther 18: 25–34, 2002. doi: 10.1089/108076802317233180. [ DOI ] [ PubMed ] [ Google Scholar ] 440. Halili L, Singh MS, Fujii N, Alexander LM, Kenny GP. Endothelin-1 modulates methacholine-induced cutaneous vasodilatation but not sweating in young human skin. J Physiol 594: 3439–52, 2016. doi: 10.1113/JP271735. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 441. Lang CC, Stein CM, Brown RM, Deegan R, Nelson R, He HB, Wood M, Wood AJ. Attenuation of isoproterenol-mediated vasodilatation in blacks. N Engl J Med 333: 155–60, 1995. doi: 10.1056/NEJM199507203330304. [ DOI ] [ PubMed ] [ Google Scholar ] 442. Eskildsen MP, Hansen PBL, Stubbe J, Toft A, Walter S, Marcussen N, Rasmussen LM, Vanhoutte PM, Jensen BL. Prostaglandin I2 and prostaglandin E2 modulate human intrarenal artery contractility through prostaglandin E2-EP4, prostacyclin-IP, and thromboxane A2-TP receptors. Hypertension 64: 551–6, 2014. doi: 10.1161/HYPERTENSIONAHA.113.03051. [ DOI ] [ PubMed ] [ Google Scholar ] 443. Lee CR, Imig JD, Edin ML, Foley J, DeGraff LM, Bradbury JA, Graves JP, Lih FB, Clark J, Myers P, Perrow AL, Lepp AN, Kannon MA, Ronnekleiv OK, Alkayed NJ, Falck JR, Tomer KB, Zeldin DC. Endothelial expression of human cytochrome P450 epoxygenases lowers blood pressure and attenuates hypertension-induced renal injury in mice. FASEB J 24: 3770–81, 2010. doi: 10.1096/fj.10-160119. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 444. Jackson-Weaver O, Osmond JM, Riddle MA, Naik JS, Gonzalez Bosc L V, Walker BR, Kanagy NL. Hydrogen sulfide dilates rat mesenteric arteries by activating endothelial large-conductance Ca 2+ -activated K + channels and smooth muscle Ca 2+ sparks. Am J Physiol Heart Circ Physiol 304: H1446–54, 2013. doi: 10.1152/ajpheart.00506.2012. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 445. Naik JS, Osmond JM, Walker BR, Kanagy NL. Hydrogen sulfide-induced vasodilation mediated by endothelial TRPV4 channels. Am J Physiol Heart Circ Physiol 311: H1437–H1444, 2016. doi: 10.1152/ajpheart.00465.2016. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 446. Shen J-K, Zhang H-T. Function and structure of bradykinin receptor 2 for drug discovery. Acta Pharmacol Sin 44: 489–498, 2023. doi: 10.1038/s41401-022-00982-8. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 447. Rabelo ER, Rohde LE, Schaan BD, Rubira MC, Ruschel KB, Plentz RDM, Consolim-Colombo FM, Irigoyen MC, Moreno Junior H. Bradykinin or acetylcholine as vasodilators to test endothelial venous function in healthy subjects. Clinics (Sao Paulo) 63: 677–82, 2008. doi: 10.1590/s1807-59322008000500017. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 448. Ceravolo GS, Fernandes L, Munhoz CD, Fernandes DC, Tostes RCA, Laurindo FRM, Scavone C, Fortes ZB, Carvalho MHC. Angiotensin II chronic infusion induces B1 receptor expression in aorta of rats. Hypertension 50: 756–61, 2007. doi: 10.1161/HYPERTENSIONAHA.107.094706. [ DOI ] [ PubMed ] [ Google Scholar ] 449. Dachman WD, Bedarida G, Blaschke TF, Hoffman BB. Histamine-induced venodilation in human beings involves both H1 and H2 receptor subtypes. J Allergy Clin Immunol 93: 606–14, 1994. doi: 10.1016/s0091-6749(94)70072-9. [ DOI ] [ PubMed ] [ Google Scholar ] 450. Haugen G, Mellembakken J, Stray-Pedersen S. Characterization of the vasodilatatory response to serotonin in human umbilical arteries perfused in vitro. The influence of the endothelium. Early Hum Dev 47: 185–93, 1997. doi: 10.1016/s0378-3782(96)01778-1. [ DOI ] [ PubMed ] [ Google Scholar ] 451. Bruning TA, Chang PC, Blauw GJ, Vermeij P, van Zwieten PA. Serotonin-induced vasodilatation in the human forearm is mediated by the “nitric oxide-pathway”: no evidence for involvement of the 5-HT3-receptor. J Cardiovasc Pharmacol 22: 44–51, 1993. doi: 10.1097/00005344-199307000-00008. [ DOI ] [ PubMed ] [ Google Scholar ] 452. Rosenblum WI, McDonald M, Wormley B. Calcium ionophore and acetylcholine dilate arterioles on the mouse brain by different mechanisms. Stroke 20: 1391–5, 1989. doi: 10.1161/01.str.20.10.1391. [ DOI ] [ PubMed ] [ Google Scholar ] 453. Gross DR, Fiscus RR, Arden WA, Maley RH, Lanzo S, Salley RK. Substance P induces biphasic endothelium-dependent relaxations in pig and rabbit carotid arteries. Neuropeptides 26: 329–41, 1994. doi: 10.1016/0143-4179(94)90118-x. [ DOI ] [ PubMed ] [ Google Scholar ] 454. Bode-Böger SM, Böger RH, Galland A, Tsikas D, Frölich JC. L-arginine-induced vasodilation in healthy humans: pharmacokinetic-pharmacodynamic relationship. Br J Clin Pharmacol 46: 489–97, 1998. doi: 10.1046/j.1365-2125.1998.00803.x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 455. Layland J, Carrick D, Lee M, Oldroyd K, Berry C. Adenosine: physiology, pharmacology, and clinical applications. JACC Cardiovasc Interv 7: 581–91, 2014. doi: 10.1016/j.jcin.2014.02.009. [ DOI ] [ PubMed ] [ Google Scholar ] 456. PAGE IH, CORCORAN AC, DUSTAN HP, KOPPANYI T. Cardiovascular actions of sodium nitroprusside in animals and hypertensive patients. Circulation 11: 188–98, 1955. doi: 10.1161/01.cir.11.2.188. [ DOI ] [ PubMed ] [ Google Scholar ] 457. Haghbin N, Richter DM, Kharche S, Kim MSM, Welsh DG. Functional bias of contractile control in mouse resistance arteries. Sci Rep 14: 24940, 2024. doi: 10.1038/s41598-024-75838-8. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 458. McKenzie C, MacDonald A, Shaw AM. Mechanisms of U46619-induced contraction of rat pulmonary arteries in the presence and absence of the endothelium. Br J Pharmacol 157: 581–96, 2009. doi: 10.1111/j.1476-5381.2008.00084.x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 459. Nelson MT, Standen NB, Brayden JE, Worley JF. Noradrenaline contracts arteries by activating voltage-dependent calcium channels. Nature 336: 382–5, 1988. doi: 10.1038/336382a0. [ DOI ] [ PubMed ] [ Google Scholar ] 460. Fransen P, Van Hove CE, Leloup AJA, Martinet W, De Meyer GRY, Lemmens K, Bult H, Schrijvers DM. Dissecting out the complex Ca2+-mediated phenylephrine-induced contractions of mouse aortic segments. PLoS One 10: e0121634, 2015. doi: 10.1371/journal.pone.0121634. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 461. Kovács A, Hársing LG, Szénási G. Vasoconstrictor 5-HT receptors in the smooth muscle of the rat middle cerebral artery. Eur J Pharmacol 689: 160–4, 2012. doi: 10.1016/j.ejphar.2012.05.031. [ DOI ] [ PubMed ] [ Google Scholar ] 462. Vanhoutte PM. Cardiovascular effects of serotonin. J Cardiovasc Pharmacol 10 Suppl 3: S8–11, 1987. [ PubMed ] [ Google Scholar ] 463. Montezano AC, Nguyen Dinh Cat A, Rios FJ, Touyz RM. Angiotensin II and vascular injury. Curr Hypertens Rep 16: 431, 2014. doi: 10.1007/s11906-014-0431-2. [ DOI ] [ PubMed ] [ Google Scholar ] 464. Griendling KK, Ushio-Fukai M, Lassègue B, Alexander RW. Angiotensin II signaling in vascular smooth muscle. New concepts. Hypertension 29: 366–73, 1997. doi: 10.1161/01.hyp.29.1.366. [ DOI ] [ PubMed ] [ Google Scholar ] 465. Tostes RC, Fortes ZB, Callera GE, Montezano AC, Touyz RM, Webb RC, Carvalho MHC. Endothelin, sex and hypertension. Clin Sci (Lond) 114: 85–97, 2008. doi: 10.1042/CS20070169. [ DOI ] [ PubMed ] [ Google Scholar ] 466. Nishiyama SK, Zhao J, Wray DW, Richardson RS. Vascular function and endothelin-1: tipping the balance between vasodilation and vasoconstriction. J Appl Physiol (1985) 122: 354–360, 2017. doi: 10.1152/japplphysiol.00772.2016. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 467. Whittle BJ, Oren-Wolman N, Guth PH. Gastric vasoconstrictor actions of leukotriene C4, PGF2 alpha, and thromboxane mimetic U-46619 on rat submucosal microcirculation in vivo. Am J Physiol 248: G580–6, 1985. doi: 10.1152/ajpgi.1985.248.5.G580. [ DOI ] [ PubMed ] [ Google Scholar ] 468. Davel AP, Lu Q, Moss ME, Rao S, Anwar IJ, DuPont JJ, Jaffe IZ. Sex-Specific Mechanisms of Resistance Vessel Endothelial Dysfunction Induced by Cardiometabolic Risk Factors. J Am Heart Assoc 7, 2018. doi: 10.1161/JAHA.117.007675. [ DOI ] [ Google Scholar ] 469. Konishi M, Su C. Role of endothelium in dilator responses of spontaneously hypertensive rat arteries. Hypertension 5: 881–6, 1983. doi: 10.1161/01.hyp.5.6.881. [ DOI ] [ PubMed ] [ Google Scholar ] 470. Lüscher TF, Vanhoutte PM. Endothelium-dependent responses to platelets and serotonin in spontaneously hypertensive rats. Hypertension 8: II55–60, 1986. doi: 10.1161/01.hyp.8.6_pt_2.ii55. [ DOI ] [ PubMed ] [ Google Scholar ] 471. Edwards JM, McCarthy CG, Wenceslau CF. The Obligatory Role of the Acetylcholine-Induced Endothelium-Dependent Contraction in Hypertension: Can Arachidonic Acid Resolve this Inflammation? Curr Pharm Des 26: 3723–3732, 2020. doi: 10.2174/1381612826666200417150121. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 472. Deng LY, Li JS, Schiffrin EL. Endothelium-dependent relaxation of small arteries from essential hypertensive patients: mechanisms and comparison with normotensive subjects and with responses of vessels from spontaneously hypertensive rats. Clin Sci (Lond) 88: 611–22, 1995. doi: 10.1042/cs0880611. [ DOI ] [ PubMed ] [ Google Scholar ] 473. Davel AP, Wenceslau CF, Akamine EH, Xavier FE, Couto GK, Oliveira HT, Rossoni LV. Endothelial dysfunction in cardiovascular and endocrine-metabolic diseases: an update. Brazilian journal of medical and biological research = Revista brasileira de pesquisas medicas e biologica 44: 920–32, 2011. doi: 10.1590/s0100-879x2011007500104. [ DOI ] [ Google Scholar ] 474. Xavier FE, Aras-López R, Arroyo-Villa I, del Campo L, Salaices M, Rossoni LV, Ferrer M, Balfagón G. Aldosterone induces endothelial dysfunction in resistance arteries from normotensive and hypertensive rats by increasing thromboxane A2 and prostacyclin. Br J Pharmacol 154: 1225–35, 2008. doi: 10.1038/bjp.2008.200. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 475. McCarthy CG, Wenceslau CF, Calmasini FB, Klee NS, Brands MW, Joe B, Webb RC. Reconstitution of autophagy ameliorates vascular function and arterial stiffening in spontaneously hypertensive rats. Am J Physiol Heart Circ Physiol 317: H1013–H1027, 2019. doi: 10.1152/ajpheart.00227.2019. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 476. Ge T, Hughes H, Junquero DC, Wu KK, Vanhoutte PM, Boulanger CM. Endothelium-dependent contractions are associated with both augmented expression of prostaglandin H synthase-1 and hypersensitivity to prostaglandin H2 in the SHR aorta. Circ Res 76: 1003–10, 1995. doi: 10.1161/01.res.76.6.1003. [ DOI ] [ PubMed ] [ Google Scholar ] 477. Hartman RJG, Kapteijn DMC, Haitjema S, Bekker MN, Mokry M, Pasterkamp G, Civelek M, den Ruijter HM. Intrinsic transcriptomic sex differences in human endothelial cells at birth and in adults are associated with coronary artery disease targets. Sci Rep 10: 12367, 2020. doi: 10.1038/s41598-020-69451-8. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 478. Orshal JM, Khalil RA. Gender, sex hormones, and vascular tone. Am J Physiol Regul Integr Comp Physiol 286: R233–49, 2004. doi: 10.1152/ajpregu.00338.2003. [ DOI ] [ PubMed ] [ Google Scholar ] 479. Smiley DA, Khalil RA. Estrogenic compounds, estrogen receptors and vascular cell signaling in the aging blood vessels. Curr Med Chem 16: 1863–87, 2009. doi: 10.2174/092986709788186093. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 480. Wong PS, Roberts RE, Randall MD. Sex differences in endothelial function in porcine coronary arteries: a role for H2O2 and gap junctions? Br J Pharmacol 171: 2751–66, 2014. doi: 10.1111/bph.12595. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 481. Marcondes FK, Bianchi FJ, Tanno AP. Determination of the estrous cycle phases of rats: some helpful considerations. Braz J Biol 62: 609–14, 2002. doi: 10.1590/s1519-69842002000400008. [ DOI ] [ PubMed ] [ Google Scholar ] 482. Iwamoto E, Sakamoto R, Tsuchida W, Yamazaki K, Kamoda T, Neki T, Katayose M, Casey DP. Effects of menstrual cycle and menopause on internal carotid artery shear-mediated dilation in women. Am J Physiol Heart Circ Physiol 320: H679–H689, 2021. doi: 10.1152/ajpheart.00810.2020. [ DOI ] [ PubMed ] [ Google Scholar ] 483. Costa TJ, Ceravolo GS, dos Santos RA, de Oliveira MA, Araújo PX, Giaquinto LR, Tostes RC, Akamine EH, Fortes ZB, Dantas AP, Carvalho MHC. Association of testosterone with estrogen abolishes the beneficial effects of estrogen treatment by increasing ROS generation in aorta endothelial cells. Am J Physiol Heart Circ Physiol 308: H723–32, 2015. doi: 10.1152/ajpheart.00681.2014. [ DOI ] [ PubMed ] [ Google Scholar ] 484. Dantas AP, Scivoletto R, Fortes ZB, Nigro D, Carvalho MH. Influence of female sex hormones on endothelium-derived vasoconstrictor prostanoid generation in microvessels of spontaneously hypertensive rats. Hypertension 34: 914–9, 1999. doi: 10.1161/01.hyp.34.4.914. [ DOI ] [ PubMed ] [ Google Scholar ] 485. Asunción-Alvarez D, Palacios J, Ybañez-Julca RO, Rodriguez-Silva CN, Nwokocha C, Cifuentes F, Greensmith DJ. Calcium signaling in endothelial and vascular smooth muscle cells: sex differences and the influence of estrogens and androgens. Am J Physiol Heart Circ Physiol 326: H950–H970, 2024. doi: 10.1152/ajpheart.00600.2023. [ DOI ] [ PubMed ] [ Google Scholar ] 486. Cattaneo MG, Vanetti C, Decimo I, Di Chio M, Martano G, Garrone G, Bifari F, Vicentini LM. Sex-specific eNOS activity and function in human endothelial cells. Sci Rep 7: 9612, 2017. doi: 10.1038/s41598-017-10139-x. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 487. Simoncini T, Maffei S, Basta G, Barsacchi G, Genazzani AR, Liao JK, De Caterina R. Estrogens and glucocorticoids inhibit endothelial vascular cell adhesion molecule-1 expression by different transcriptional mechanisms. Circ Res 87: 19–25, 2000. doi: 10.1161/01.res.87.1.19. [ DOI ] [ PubMed ] [ Google Scholar ] 488. Lorenz M, Koschate J, Kaufmann K, Kreye C, Mertens M, Kuebler WM, Baumann G, Gossing G, Marki A, Zakrzewicz A, Miéville C, Benn A, Horbelt D, Wratil PR, Stangl K, Stangl V. Does cellular sex matter? Dimorphic transcriptional differences between female and male endothelial cells. Atherosclerosis 240: 61–72, 2015. doi: 10.1016/j.atherosclerosis.2015.02.018. [ DOI ] [ PubMed ] [ Google Scholar ] 489. Weber CM, Clyne AM. Sex differences in the blood-brain barrier and neurodegenerative diseases. APL Bioeng 5: 011509, 2021. doi: 10.1063/5.0035610. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 490. Davis MJ. An improved, computer-based method to automatically track internal and external diameter of isolated microvessels. Microcirculation 12: 361–72, 2005. doi: 10.1080/10739680590934772. [ DOI ] [ PubMed ] [ Google Scholar ] 491. Lawton PF, Lee MD, Saunter CD, Girkin JM, McCarron JG, Wilson C. VasoTracker, a Low-Cost and Open Source Pressure Myograph System for Vascular Physiology. Front Physiol 10: 99, 2019. doi: 10.3389/fphys.2019.00099. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 492. Lee MD, Osborne C, Stevenson R, MacDonald A, Ebner G, Jeffrey DA, Macdonald MA, Zhang X, Buckley C, Dabertrand F, Machin DR, Au J, Harraz OF, Tykocki N, McCarron JG, Wilson C. VasoTracker 2: An Open-source Platform for Quantitative Analysis of Vascular Reactivity and Function. 2025. [ Google Scholar ] 493. Edelstein A, Amodaj N, Hoover K, Vale R, Stuurman N. Computer control of microscopes using μManager. Curr Protoc Mol Biol Chapter 14: Unit14.20, 2010. doi: 10.1002/0471142727.mb1420s92. [ DOI ] [ Google Scholar ] 494. Lee MD, Wilson C, Saunter CD, Kennedy C, Girkin JM, McCarron JG. Spatially structured cell populations process multiple sensory signals in parallel in intact vascular endothelium. Sci Signal 11, 2018. doi: 10.1126/scisignal.aar4411. [ DOI ] [ Google Scholar ] 495. Buckley C, Lee MD, Zhang X, Wilson C, McCarron JG. Signalling switches maintain intercellular communication in the vascular endothelium. Br J Pharmacol 181: 2810–2832, 2024. doi: 10.1111/bph.16366. [ DOI ] [ PubMed ] [ Google Scholar ] 496. Dempster J, Wokosin D, McCloskey K, Girkin J, Gurney A. WinFluor-an integrated system for the simultaneous recording of cell fluorescence images and electrophysiological signals on a single computer system. In: Br. J. Pharmacol 2002. [ Google Scholar ] 497. Schneider CA, Rasband WS, Eliceiri KW. NIH Image to ImageJ: 25 years of image analysis. Nat Methods 9: 671–5, 2012. doi: 10.1038/nmeth.2089. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 498. Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch T, Preibisch S, Rueden C, Saalfeld S, Schmid B, Tinevez J-Y, White DJ, Hartenstein V, Eliceiri K, Tomancak P, Cardona A. Fiji: an open-source platform for biological-image analysis. Nat Methods 9: 676–82, 2012. doi: 10.1038/nmeth.2019. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 499. Knighten JM, Aziz T, Pleshinger DJ, Annamdevula N, Rich TC, Taylor MS, Andrews JF, Macarilla CT, Francis CM. Algorithm for biological second messenger analysis with dynamic regions of interest. PLoS One 18: e0284394, 2023. doi: 10.1371/journal.pone.0284394. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 500. Wilson C, Lee MD, McCarron JG. Acetylcholine released by endothelial cells facilitates flow-mediated dilatation. J Physiol 594: 7267–7307, 2016. doi: 10.1113/JP272927. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 501. Lee MD, Buckley C, Zhang X, Louhivuori L, Uhlén P, Wilson C, McCarron JG. Small-world connectivity dictates collective endothelial cell signaling. Proc Natl Acad Sci U S A 119: e2118927119, 2022. doi: 10.1073/pnas.2118927119. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 502. Picht E, Zima AV, Blatter LA, Bers DM. SparkMaster: automated calcium spark analysis with ImageJ. Am J Physiol Cell Physiol 293: C1073–81, 2007. doi: 10.1152/ajpcell.00586.2006. [ DOI ] [ PubMed ] [ Google Scholar ] 503. Tomek J, Nieves-Cintron M, Navedo MF, Ko CY, Bers DM. SparkMaster 2: A New Software for Automatic Analysis of Calcium Spark Data. Circ Res 133: 450–462, 2023. doi: 10.1161/CIRCRESAHA.123.322847. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 504. Ellefsen KL, Lock JT, Settle B, Karsten CA, Parker I. Applications of FLIKA, a Python-based image processing and analysis platform, for studying local events of cellular calcium signaling. Biochim Biophys Acta Mol Cell Res 1866: 1171–1179, 2019. doi: 10.1016/j.bbamcr.2018.11.012. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 505. Wilson C, Zhang X, Buckley C, Heathcote HR, Lee MD, McCarron JG. Increased Vascular Contractility in Hypertension Results From Impaired Endothelial Calcium Signaling. Hypertension 74: 1200–1214, 2019. doi: 10.1161/HYPERTENSIONAHA.119.13791. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 506. Giovannucci A, Friedrich J, Gunn P, Kalfon J, Brown BL, Koay SA, Taxidis J, Najafi F, Gauthier JL, Zhou P, Khakh BS, Tank DW, Chklovskii DB, Pnevmatikakis EA. CaImAn an open source tool for scalable calcium imaging data analysis. Elife 8, 2019. doi: 10.7554/eLife.38173. [ DOI ] [ Google Scholar ] 507. Napari contributors. napari: a multi-dimensional image viewer for Python [software]. Zenodo: 2019. [ Google Scholar ] 508. Takahashi K, Yamanaka S. Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 126: 663–76, 2006. doi: 10.1016/j.cell.2006.07.024. [ DOI ] [ PubMed ] [ Google Scholar ] 509. Takahashi K, Tanabe K, Ohnuki M, Narita M, Ichisaka T, Tomoda K, Yamanaka S. Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 131: 861–72, 2007. doi: 10.1016/j.cell.2007.11.019. [ DOI ] [ PubMed ] [ Google Scholar ] 510. Nguyen V, Gao C, Hochman ML, Kravitz J, Chen EH, Friedman HI, Wenceslau CF, Chen D, Wang Y, Nelson JS, Jegga AG, Tan W. Endothelial cells differentiated from patient dermal fibroblast-derived induced pluripotent stem cells resemble vascular malformations of port-wine birthmark. Br J Dermatol 189: 780–783, 2023. doi: 10.1093/bjd/ljad330. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 511. Sahara M, Hansson EM, Wernet O, Lui KO, Später D, Chien KR. Manipulation of a VEGF-Notch signaling circuit drives formation of functional vascular endothelial progenitors from human pluripotent stem cells. Cell Res 25: 148, 2015. doi: 10.1038/cr.2015.2. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 512. James D, Nam H, Seandel M, Nolan D, Janovitz T, Tomishima M, Studer L, Lee G, Lyden D, Benezra R, Zaninovic N, Rosenwaks Z, Rabbany SY, Rafii S. Expansion and maintenance of human embryonic stem cell-derived endothelial cells by TGFbeta inhibition is Id1 dependent. Nat Biotechnol 28: 161–6, 2010. doi: 10.1038/nbt.1605. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 513. Bao X, Lian X, Palecek SP. Directed Endothelial Progenitor Differentiation from Human Pluripotent Stem Cells Via Wnt Activation Under Defined Conditions. Methods Mol Biol 1481: 183–96, 2016. doi: 10.1007/978-1-4939-6393-5_17. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 514. Ikuno T, Masumoto H, Yamamizu K, Yoshioka M, Minakata K, Ikeda T, Sakata R, Yamashita JK. Efficient and robust differentiation of endothelial cells from human induced pluripotent stem cells via lineage control with VEGF and cyclic AMP. PLoS One 12: e0173271, 2017. doi: 10.1371/journal.pone.0173271. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 515. Glaser DE, Turner WS, Madfis N, Wong L, Zamora J, White N, Reyes S, Burns AB, Gopinathan A, McCloskey KE. Multifactorial Optimizations for Directing Endothelial Fate from Stem Cells. PLoS One 11: e0166663, 2016. doi: 10.1371/journal.pone.0166663. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 516. Cochrane A, Kelaini S, Tsifaki M, Bojdo J, Vilà-González M, Drehmer D, Caines R, Magee C, Eleftheriadou M, Hu Y, Grieve D, Stitt AW, Zeng L, Xu Q, Margariti A. Quaking Is a Key Regulator of Endothelial Cell Differentiation, Neovascularization, and Angiogenesis. Stem Cells 35: 952–966, 2017. doi: 10.1002/stem.2594. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 517. Liang J, Huang W, Cai W, Wang L, Guo L, Paul C, Yu X-Y, Wang Y. Inhibition of microRNA-495 Enhances Therapeutic Angiogenesis of Human Induced Pluripotent Stem Cells. Stem Cells 35: 337–350, 2017. doi: 10.1002/stem.2477. [ DOI ] [ PubMed ] [ Google Scholar ] 518. Noh KM, Park S-J, Moon S-H, Jung SY. Extracellular matrix cues regulate the differentiation of pluripotent stem cell-derived endothelial cells. Front Cardiovasc Med 10: 1169331, 2023. doi: 10.3389/fcvm.2023.1169331. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 519. Ohtani-Kaneko R, Sato K, Tsutiya A, Nakagawa Y, Hashizume K, Tazawa H. Characterisation of human induced pluripotent stem cell-derived endothelial cells under shear stress using an easy-to-use microfluidic cell culture system. Biomed Microdevices 19: 91, 2017. doi: 10.1007/s10544-017-0229-5. [ DOI ] [ PubMed ] [ Google Scholar ] 520. Rufaihah AJ, Huang NF, Kim J, Herold J, Volz KS, Park TS, Lee JC, Zambidis ET, Reijo-Pera R, Cooke JP. Human induced pluripotent stem cell-derived endothelial cells exhibit functional heterogeneity. Am J Transl Res 5: 21–35, 2013. [ PMC free article ] [ PubMed ] [ Google Scholar ] 521. Zhang J, Chu L-F, Hou Z, Schwartz MP, Hacker T, Vickerman V, Swanson S, Leng N, Nguyen BK, Elwell A, Bolin J, Brown ME, Stewart R, Burlingham WJ, Murphy WL, Thomson JA. Functional characterization of human pluripotent stem cell-derived arterial endothelial cells. Proc Natl Acad Sci U S A 114: E6072–E6078, 2017. doi: 10.1073/pnas.1702295114. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 522. Sivarapatna A, Ghaedi M, Le AV, Mendez JJ, Qyang Y, Niklason LE. Arterial specification of endothelial cells derived from human induced pluripotent stem cells in a biomimetic flow bioreactor. Biomaterials 53: 621–33, 2015. doi: 10.1016/j.biomaterials.2015.02.121. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 523. Ang LT, Nguyen AT, Liu KJ, Chen A, Xiong X, Curtis M, Martin RM, Raftry BC, Ng CY, Vogel U, Lander A, Lesch BJ, Fowler JL, Holman AR, Chai T, Vijayakumar S, Suchy FP, Nishimura T, Bhadury J, Porteus MH, Nakauchi H, Cheung C, George SC, Red-Horse K, Prescott JB, Loh KM. Generating human artery and vein cells from pluripotent stem cells highlights the arterial tropism of Nipah and Hendra viruses. Cell 185: 2523–2541.e30, 2022. doi: 10.1016/j.cell.2022.05.024. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 524. Lee S-J, Park C, Lee JY, Kim S, Kwon PJ, Kim W, Jeon YH, Lee E, Yoon Y. Generation of pure lymphatic endothelial cells from human pluripotent stem cells and their therapeutic effects on wound repair. Sci Rep 5: 11019, 2015. doi: 10.1038/srep11019. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 525. Linville RM, DeStefano JG, Sklar MB, Xu Z, Farrell AM, Bogorad MI, Chu C, Walczak P, Cheng L, Mahairaki V, Whartenby KA, Calabresi PA, Searson PC. Human iPSC-derived blood-brain barrier microvessels: validation of barrier function and endothelial cell behavior. Biomaterials 190–191: 24–37, 2019. doi: 10.1016/j.biomaterials.2018.10.023. [ DOI ] [ Google Scholar ] 526. Stebbins MJ, Wilson HK, Canfield SG, Qian T, Palecek SP, Shusta EV. Differentiation and characterization of human pluripotent stem cell-derived brain microvascular endothelial cells. Methods 101: 93–102, 2016. doi: 10.1016/j.ymeth.2015.10.016. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 527. Lian X, Bao X, Al-Ahmad A, Liu J, Wu Y, Dong W, Dunn KK, Shusta EV, Palecek SP. Efficient differentiation of human pluripotent stem cells to endothelial progenitors via small-molecule activation of WNT signaling. Stem Cell Reports 3: 804–16, 2014. doi: 10.1016/j.stemcr.2014.09.005. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 528. So S, Park Y, Kang SS, Han J, Sunwoo JH, Lee W, Kim J, Ye EA, Kim JY, Tchah H, Kang E, Lee H. Therapeutic Potency of Induced Pluripotent Stem-Cell-Derived Corneal Endothelial-like Cells for Corneal Endothelial Dysfunction. Int J Mol Sci 24, 2022. doi: 10.3390/ijms24010701. [ DOI ] [ Google Scholar ] 529. Mulfaul K, Giacalone JC, Voigt AP, Riker MJ, Ochoa D, Han IC, Stone EM, Mullins RF, Tucker BA. Stepwise differentiation and functional characterization of human induced pluripotent stem cell-derived choroidal endothelial cells. Stem Cell Res Ther 11: 409, 2020. doi: 10.1186/s13287-020-01903-4. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 530. Prasain N, Lee MR, Vemula S, Meador JL, Yoshimoto M, Ferkowicz MJ, Fett A, Gupta M, Rapp BM, Saadatzadeh MR, Ginsberg M, Elemento O, Lee Y, Voytik-Harbin SL, Chung HM, Hong KS, Reid E, O’Neill CL, Medina RJ, Stitt AW, Murphy MP, Rafii S, Broxmeyer HE, Yoder MC. Differentiation of human pluripotent stem cells to cells similar to cord-blood endothelial colony-forming cells. Nat Biotechnol 32: 1151–1157, 2014. doi: 10.1038/nbt.3048. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 531. Romeo SG, Secco I, Schneider E, Reumiller CM, Santos CXC, Zoccarato A, Musale V, Pooni A, Yin X, Theofilatos K, Trevelin SC, Zeng L, Mann GE, Pathak V, Harkin K, Stitt AW, Medina RJ, Margariti A, Mayr M, Shah AM, Giacca M, Zampetaki A. Human blood vessel organoids reveal a critical role for CTGF in maintaining microvascular integrity. Nat Commun 14: 5552, 2023. doi: 10.1038/s41467-023-41326-2. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 532. Matrone G, Thandavarayan RA, Walther BK, Meng S, Mojiri A, Cooke JP. Dysfunction of iPSC-derived endothelial cells in human Hutchinson-Gilford progeria syndrome. Cell Cycle 18: 2495–2508, 2019. doi: 10.1080/15384101.2019.1651587. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 533. Dao L, You Z, Lu L, Xu T, Sarkar AK, Zhu H, Liu M, Calandrelli R, Yoshida G, Lin P, Miao Y, Mierke S, Kalva S, Zhu H, Gu M, Vadivelu S, Zhong S, Huang LF, Guo Z. Modeling blood-brain barrier formation and cerebral cavernous malformations in human PSC-derived organoids. Cell Stem Cell 31: 818–833.e11, 2024. doi: 10.1016/j.stem.2024.04.019. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 534. Zhou D, Tan Y, Liu X, Tang L, Wang H, Shen J, Wang W, Zhuang L, Tao J, Su J, Gong T, Liu X, Liang P, Yu F, Zhao M. Patient-specific iPSC-derived endothelial cells reveal aberrant p38 MAPK signaling in atypical hemolytic uremic syndrome. Stem Cell Reports 16: 2305–2319, 2021. doi: 10.1016/j.stemcr.2021.07.011. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 535. Zakharova IS, Shevchenko AI, Arssan MA, Sleptcov AA, Nazarenko MS, Zarubin AA, Zheltysheva NV, Shevchenko VA, Tmoyan NA, Saaya SB, Ezhov MV, Kukharchuk VV, Parfyonova YV, Zakian SM. iPSC-Derived Endothelial Cells Reveal LDLR Dysfunction and Dysregulated Gene Expression Profiles in Familial Hypercholesterolemia. Int J Mol Sci 25, 2024. doi: 10.3390/ijms25020689. [ DOI ] [ Google Scholar ] 536. DeStefano JG, Xu ZS, Williams AJ, Yimam N, Searson PC. Effect of shear stress on iPSC-derived human brain microvascular endothelial cells (dhBMECs). Fluids Barriers CNS 14: 20, 2017. doi: 10.1186/s12987-017-0068-z. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 537. Park S-W, Jun Koh Y, Jeon J, Cho Y-H, Jang M-J, Kang Y, Kim M-J, Choi C, Sook Cho Y, Chung H-M, Koh GY, Han Y-M. Efficient differentiation of human pluripotent stem cells into functional CD34+ progenitor cells by combined modulation of the MEK/ERK and BMP4 signaling pathways. Blood 116: 5762–72, 2010. doi: 10.1182/blood-2010-04-280719. [ DOI ] [ PubMed ] [ Google Scholar ] 538. Zhang F, Wang L, Li Y, Liu W, Duan F, Huang R, Chen X, Chang SC- N, Du Y, Na J. Optimizing mesoderm progenitor selection and three-dimensional microniche culture allows highly efficient endothelial differentiation and ischemic tissue repair from human pluripotent stem cells. Stem Cell Res Ther 8: 6, 2017. doi: 10.1186/s13287-016-0455-4. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 539. Nishihara H, Gastfriend BD, Soldati S, Perriot S, Mathias A, Sano Y, Shimizu F, Gosselet F, Kanda T, Palecek SP, Du Pasquier R, Shusta EV, Engelhardt B. Advancing human induced pluripotent stem cell-derived blood-brain barrier models for studying immune cell interactions. FASEB J 34: 16693–16715, 2020. doi: 10.1096/fj.202001507RR. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 540. Straessler ET, Kiamehr M, Aalto-Setala K, Kraenkel NK. Straessler ET, Kiamehr M, Aalto-Setala K, Kraenkel NK, Landmesser UL. iPSC-derived endothelial cells reflect accelerated senescence and increased inflammatory response in a CVD-risk stratified manner. European Heart Journal, Volume 41, Issue Supplement_2, November 2020. In: iPSC-derived endothelial cells reflect accelerated senescence and increased inflammatory response in a CVD-risk stratified manner. 2020. [ Google Scholar ] 541. Shakeri A, Wang Y, Zhao Y, Landau S, Perera K, Lee J, Radisic M. Engineering Organ-on-a-Chip Systems for Vascular Diseases. Arterioscler Thromb Vasc Biol 43: 2241–2255, 2023. doi: 10.1161/ATVBAHA.123.318233. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 542. Shiwarski DJ, Hudson AR, Tashman JW, Bakirci E, Moss S, Coffin BD, Feinberg AW. 3D bioprinting of collagen-based high-resolution internally perfusable scaffolds for engineering fully biologic tissue systems. Sci Adv 11: eadu5905, 2025. doi: 10.1126/sciadv.adu5905. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 543. Venugopal Menon N, Tay HM, Pang KT, Dalan R, Wong SC, Wang X, Li KHH, Hou HW. A tunable microfluidic 3D stenosis model to study leukocyte-endothelial interactions in atherosclerosis. APL Bioeng 2: 016103, 2018. doi: 10.1063/1.4993762. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 544. Yang Q, Langston JC, Prosniak R, Pettigrew S, Zhao H, Perez E, Edelmann H, Mansoor N, Merali C, Merali S, Marchetti N, Prabhakarpandian B, Kiani MF, Kilpatrick LE. Distinct functional neutrophil phenotypes in sepsis patients correlate with disease severity. Front Immunol 15: 1341752, 2024. doi: 10.3389/fimmu.2024.1341752. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 545. Ng WH, Varghese B, Ren X. Understanding and Engineering the Pulmonary Vasculature. Adv Exp Med Biol 1413: 247–264, 2023. doi: 10.1007/978-3-031-26625-6_12. [ DOI ] [ PubMed ] [ Google Scholar ] 546. Yuan Y, Leiby KL, Greaney AM, Raredon MSB, Qian H, Schupp JC, Engler AJ, Baevova P, Adams TS, Kural MH, Wang J, Obata T, Yoder MC, Kaminski N, Niklason LE. A Pulmonary Vascular Model From Endothelialized Whole Organ Scaffolds. Front Bioeng Biotechnol 9: 760309, 2021. doi: 10.3389/fbioe.2021.760309. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 547. Wang Z, Zhang P. Novel imaging modalities for the identification of vulnerable plaques. Front Cardiovasc Med 11: 1450252, 2024. doi: 10.3389/fcvm.2024.1450252. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 548. Colcombe J, Solli E, Kaiser A, Ranadive I, Bolneni S, Berger J, Garshick M, Modi Y. The Use of Retinal Imaging Including Fundoscopy, OCT, and OCTA for Cardiovascular Risk Stratification and the Detection of Subclinical Atherosclerosis. Curr Atheroscler Rep 27: 23, 2025. doi: 10.1007/s11883-024-01268-6. [ DOI ] [ PubMed ] [ Google Scholar ] 549. Chen J, Li S, Zhou Q, Zhao X, Fan Z, Lo H, Nie L. Near-Infrared II Fluorescence Imaging Highlights Tumor Angiogenesis in Hepatocellular Carcinoma with a VEGFR-Targeted Probe. Small Methods 9: e2400904, 2025. doi: 10.1002/smtd.202400904. [ DOI ] [ PubMed ] [ Google Scholar ] 550. Yu F, Zhang Y, Sun H, Li X, Shan Y, Zheng C, Cui B, Li J, Yang Y, Yang B, Ma Y, Wang Y, Jiao L, Li X, Lu J. In Vivo Classification and Characterization of Carotid Atherosclerotic Lesions with Integrated 18F-FDG PET/MRI. Diagnostics (Basel) 14, 2024. doi: 10.3390/diagnostics14101006. [ DOI ] [ Google Scholar ] 551. Deshayes S, Ruello P, Simard C, Dupont P-A, Bauge C, Abbas A, de Boysson H, Aouba A, Manrique A. 18F-fluorodeoxyglucose PET-MR characterization of aortic inflammation in ApoE−/− mouse models of accelerated atherosclerosis: comparison of Western diet vs. uremia. Int J Cardiovasc Imaging 40: 2335–2344, 2024. doi: 10.1007/s10554-024-03238-0. [ DOI ] [ PubMed ] [ Google Scholar ] 552. Schilling K, Zhai Y, Zhou Z, Zhou B, Brown E, Zhang X. High-resolution imaging of the osteogenic and angiogenic interface at the site of murine cranial bone defect repair via multiphoton microscopy. Elife 11, 2022. doi: 10.7554/eLife.83146. [ DOI ] [ Google Scholar ] 553. Zhang D, Cleveland AH, Krimitza E, Han K, Yi C, Stout AL, Zou W, Dorsey JF, Gong Y, Fan Y. Spatial analysis of tissue immunity and vascularity by light sheet fluorescence microscopy. Nat Protoc 19: 1053–1082, 2024. doi: 10.1038/s41596-023-00941-5. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 554. Deep Ertürk A. 3D histology powered by tissue clearing, omics and AI. Nat Methods 21: 1153–1165, 2024. doi: 10.1038/s41592-024-02327-1. [ DOI ] [ PubMed ] [ Google Scholar ] 555. Huang M, Yang F, Zhang D, Lin M, Duan H, El-Mayta R, Zhang L, Qin L, Shewale SV, Pei L, Mitchell MJ, Rader DJ, Fan Y, Gong Y. Endothelial plasticity drives aberrant vascularization and impedes cardiac repair after myocardial infarction. Nature cardiovascular research 1: 372–388, 2022. doi: 10.1038/s44161-022-00047-3. [ DOI ] [ Google Scholar ] 556. Bennett HC, Zhang Q, Wu Y-T, Manjila SB, Chon U, Shin D, Vanselow DJ, Pi H-J, Drew PJ, Kim Y. Aging drives cerebrovascular network remodeling and functional changes in the mouse brain. Nat Commun 15: 6398, 2024. doi: 10.1038/s41467-024-50559-8. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 557. Kalucka J, de Rooij LPMH, Goveia J, Rohlenova K, Dumas SJ, Meta E, Conchinha NV, Taverna F, Teuwen L-A, Veys K, García-Caballero M, Khan S, Geldhof V, Sokol L, Chen R, Treps L, Borri M, de Zeeuw P, Dubois C, Karakach TK, Falkenberg KD, Parys M, Yin X, Vinckier S, Du Y, Fenton RA, Schoonjans L, Dewerchin M, Eelen G, Thienpont B, Lin L, Bolund L, Li X, Luo Y, Carmeliet P. Single-Cell Transcriptome Atlas of Murine Endothelial Cells. Cell 180: 764–779.e20, 2020. doi: 10.1016/j.cell.2020.01.015. [ DOI ] [ PubMed ] [ Google Scholar ] 558. Ma X, Chen P, Wei J, Zhang J, Chen C, Zhao H, Ferguson D, McGee AW, Dai Z, Qiu S. Protocol for Xenium spatial transcriptomics studies using fixed frozen mouse brain sections. STAR Protoc 5: 103420, 2024. doi: 10.1016/j.xpro.2024.103420. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 559. Liu B, Yi D, Xia X, Ramirez K, Zhao H, Cao Y, Tripathi A, Dong R, Gao A, Ding H, Qiu S, Kalinichenko VV, Zhao Y-Y, Fallon MB, Dai Z. General Capillary Endothelial Cells Undergo Reprogramming Into Arterial Endothelial Cells in Pulmonary Hypertension Through HIF-2α/Notch4 Pathway. Circulation 150: 414–417, 2024. doi: 10.1161/CIRCULATIONAHA.123.067981. [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] 560. Subramanian A, Narayan R, Corsello SM, Peck DD, Natoli TE, Lu X, Gould J, Davis JF, Tubelli AA, Asiedu JK, Lahr DL, Hirschman JE, Liu Z, Donahue M, Julian B, Khan M, Wadden D, Smith IC, Lam D, Liberzon A, Toder C, Bagul M, Orzechowski M, Enache OM, Piccioni F, Johnson SA, Lyons NJ, Berger AH, Shamji AF, Brooks AN, Vrcic A, Flynn C, Rosains J, Takeda DY, Hu R, Davison D, Lamb J, Ardlie K, Hogstrom L, Greenside P, Gray NS, Clemons PA, Silver S, Wu X, Zhao W-N, Read-Button W, Wu X, Haggarty SJ, Ronco LV, Boehm JS, Schreiber SL, Doench JG, Bittker JA, Root DE, Wong B, Golub TR. A Next Generation Connectivity Map: L1000 Platform and the First 1,000,000 Profiles. Cell 171: 1437–1452.e17, 2017. doi: 10.1016/j.cell.2017.10.049.</References> [ DOI ] [ PMC free article ] [ PubMed ] [ Google Scholar ] ACTIONS View on publisher site PDF (5.7 MB) Cite Collections Permalink PERMALINK Copy RESOURCES Similar articles Cited by other articles Links to NCBI Databases Cite Copy Download .nbib .nbib Format: AMA APA MLA NLM Add to Collections Create a new collection Add to an existing collection Name your collection * Choose a collection Unable to load your collection due to an error Please try again Add Cancel Follow NCBI NCBI on X (formerly known as Twitter) NCBI on Facebook NCBI on LinkedIn NCBI on GitHub NCBI RSS feed Connect with NLM NLM on X (formerly known as Twitter) NLM on Facebook NLM on YouTube National Library of Medicine 8600 Rockville Pike Bethesda, MD 20894 Web Policies FOIA HHS Vulnerability Disclosure Help Accessibility Careers NLM NIH HHS USA.gov Back to Top

Related documents

• Relapse prediction and individualized treatment-effect modeling in relapsing multiple sclerosis: a systematic review.#1016102