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Method Article

Multilevel Microdissection and Functional-Structural Profiling of Human Renal Arterial Branches

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DOI:

10.3791/68579

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September 5th, 2025

 ,  ,  ,  ,  ,  , 
1Institute of Cardiovascular Sciences, School of Basic Medical Sciences, Peking University Health Science Center; State Key Laboratory of Vascular Homeostasis and Remodeling, Peking University, 2Beijing Key Laboratory of Cardiovascular Receptors Research, 3Department of Cardiology and Institute of Vascular Medicine, Peking University Third Hospital, Peking University, 4Department of Urology, Peking University First Hospital; Institute of Urology, Peking University; Beijing Key Laboratory of Urogenital Diseases (Male) Molecular Diagnosis and Treatment Center, National Urological Cancer Center, 5Institute of Cardiovascular Diseases, First Affiliated Hospital of Dalian Medical University, 6NHC Key Laboratory of Cardiovascular Molecular Biology and Regulatory Peptides, Peking University

In This Article

Summary

This article presents a step-by-step protocol for the isolation and functional evaluation of human renal arterial branches, facilitating preclinical studies for pharmaceutical development.

Abstract

Renal vascular dysfunction plays a critical role in the pathogenesis of multiple clinical conditions, including acute kidney injury, renal ischemia, and hypertension, presenting significant challenges in clinical management and adversely affecting patient outcomes. The isolation and functional characterization of intrarenal arteries are crucial for elucidating the mechanisms underlying renal vascular dysfunction, particularly related to kidney injury, and guiding targeted therapeutic development. Despite its clinical importance, standardized approaches for isolating and functionally assessing human intrarenal arteries across different branching levels remain underdeveloped. This protocol provides a comprehensive framework for the systematic isolation and multimodal evaluation of intrarenal arterial branches, incorporating functional and structural assessments under both physiological and pathological conditions. The methodology encompasses three key components: (1) precise anatomical identification and microdissection of intrarenal arteries from donor kidneys, accompanied by Hematoxylin-Eosin (H&E) staining for structural confirmation; (2) rigorous normalization procedures in wire myography to enhance measurement reproducibility and reliability; and (3) quantitative analysis of vasomotor responses using precision wire myography techniques. Normalization is based on the muscle length-tension relationship, where incremental stretching of arterial segments establishes an optimal resting tension to maximize actin-myosin overlap, thereby eliciting peak contractile responses. In wire myography, isolated vessel segments are suspended between two parallel wires, allowing precise measurement of vascular tension. By applying rigorous normalization protocols, this technique enables reproducible and reliable quantification of vascular reactivity across diverse pathophysiological conditions and pharmacological interventions.

Introduction

The kidney is a vital organ responsible for maintaining human homeostasis through clearance of metabolic wastes, electrolyte balance, and fluid regulation - functions that demand exceptionally high blood flow. Under physiological conditions, the kidneys receive approximately 25% of the total blood volume pumped by the heart each minute (cardiac output), underscoring their substantial role in systemic blood flow and the critical importance of renal perfusion in maintaining overall physiological balance1. The renal arterial system precisely regulates this massive blood delivery to maintain GFR while adapting to systemic hemodynamic changes. Consequently, impairment of renal perfusion represents a critical pathological mechanism that contributes to reduced glomerular filtration rate (GFR) in conditions such as acute kidney injury and renal ischemia. In addition, it frequently occurs secondary to systemic disorders that compromise renal blood flow, including shock, cardiac dysfunction, and sepsis2,3,4,5. The renal artery undergoes sequential branching within the kidney, first dividing into interlobar arteries that course through the renal columns between medullary pyramids. Upon reaching the corticomedullary junction, these vessels give rise to arcuate arteries, which arch along the boundary between cortex and medulla. From these curvilinear vessels emerge interlobular arteries that penetrate radially into the cortical parenchyma, ultimately delivering blood to the glomerular capillaries via afferent arterioles to perfuse individual nephrons1,6,7. Interlobar, arcuate, and interlobular arteries constitute the key resistance vasculature in the kidney, serving as primary regulators of intrarenal hemodynamics and pressure control1,8. Precise isolation and characterization of these vessels are crucial for elucidating the fundamental mechanisms governing renal perfusion homeostasis9. Furthermore, their heterogeneous vascular reactivity under various pathophysiological conditions offers a strategic framework for developing novel vasodilatory agents with renal-protective potential10,11.

Currently, clinical imaging techniques (e.g., ultrasound) are routinely used to monitor renal blood flow changes; however, experimental methodologies in basic research for directly assessing renal vascular tension, a key determinant of renal perfusion, remain limited12,13,14,15. The wire myography system is a well-established experimental platform that measures tension changes in small biological tissues in vitro, including blood vessels, trachea16,17,18, with diameters ranging from 50 µm to 10 mm16,19,20. Although wire myography has been widely applied in animal studies, interspecies differences in vascular structure and function limit the extent to which rodent vessels can represent human vascular physiology21. Therefore, precise identification and isolation of human intrarenal arteries at various anatomical levels (interlobar, arcuate, and interlobular) are crucial for enabling comprehensive in vitro investigations into their physiological properties, as well as disease-related functional alterations and pathological injury mechanisms. When combined with pharmacological interventions using various vasoactive compounds, this system allows for the dynamic assessment and quantitative recording of vasoconstriction/vasodilation responses in isolated arterial rings, providing a powerful tool for investigating vascular reactivity under controlled experimental conditions19,22,23.

This protocol details a comprehensive methodology for the precise anatomical localization and microdissection of human intarenal artery branches (interlobar, arcuate, and interlobular arteries), followed by functional characterization using an in vitro vascular tension monitoring system to assess their differential reactivity to vasoactive drugs, thereby establishing both structural and functional distinctions between these vascular segments. By combining anatomical localization with pharmacological reactivity profiling and histopathological analysis, we establish an ex vivo platform that enables comprehensive evaluation of region-specific vascular responses to vasoactive agents.

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Protocol

The protocol and examples described here were reviewed and approved by the Urology Department of Peking University First Hospital (approval No. 2023yan500-002) and conducted in accordance with the Helsinki Declaration. All participants provided written informed consent prior to participation.

1. Solution preparation

  1. Prepare Krebs-Ringer solution (Krebs) containing 119.0 mM NaCl, 4.7 mM KCl, 2.5 mM CaCl2, 1.0 mM MgCl2, 25.0 mM NaHCO3, 1.2 mM KH2PO4, and 11.0 mM D-glucose at pH 7.4.
  2. Prepare high potassium salt solution (60K+) containing 64.0 mM NaCl, 60 mM KCl, 2.5 mM CaCl2, 1.0 mM MgCl2, 25.0 mM NaHCO3, 1.2 mM KH2PO4, and 11.0 mM D-glucose at pH 7.4.

2. Reagent preparation

NOTE: Store drugs as per the details provided in the Material Safety Data Sheet (MSDS) immediately after receiving them.

  1. Prepare the phenylephrine (Phe, 10-1 M in stock) by dissolving 167.21 mg of phenylephrine in 10 mL of dimethyl sulfoxide (DMSO). Store the stock solution at -80 °C, dilute to 10-6, 10-5, 10-4, 10-3, 10-2 M as working concentration on the day of use.
  2. Prepare the acetylcholine (ACh, 10-2 M in stock), by dissolving 18.166 mg of acetylcholine in 10 mL of dimethyl sulfoxide (DMSO). Store stock solution at -80 °C, dilute to 10-5, 10-4, 10-3, 10-2 M as working concentration on the day of use.

3. Kidney tissue collection and intrarenal artery isolation

  1. Recruit adult patients diagnosed with renal cell carcinoma, as histologically confirmed by preoperative core needle biopsy or intraoperative frozen section.
    NOTE: Patients scheduled for radical nephrectomy are specifically chosen because their surgical specimens provide access to intact intrarenal arterial segments located distal to tumor margins. These segments can be precisely dissected and isolated, allowing detailed study of human renal vascular physiology without interference from malignant tissue. Histological confirmation ensures accurate identification of carcinoma and clear delineation of tumor boundaries, facilitating the selection of normal, tumor-free arterial segments for experimental investigations.
    1. Exclude patients with significant comorbidities (hypertension, diabetes mellitus, or other systemic diseases) to avoid confounding influences on vascular function, thus enhancing the reliability and relevance of physiological assessments.
      NOTE: The samples for this experiment were collected from patients scheduled for radical nephrectomy at Peking University First Hospital.
  2. During radical nephrectomy, collect freshly excised human kidney specimens and immediately immerse them in ice-cold Krebs solution to preserve tissue viability. Only macroscopically normal-appearing regions distal to tumor margins are retained for arterial dissection (Figure 1A).
    NOTE: For optimal intrarenal artery isolation, adjacent healthy tissue should be harvested as a full-thickness corticomedullary wedge during radical nephrectomy, thereby preserving the complete arterial tree from cortical to medullary vasculature.
    1. Immediately store the excised tissue in 200 mL of cold Krebs solution. Place the container on ice and transport it in an insulated cooler bag to the laboratory within 2 h of collection. Ensure that the tissue remains fully submerged throughout transport to preserve viability and structural integrity for subsequent dissection and functional analysis.
      NOTE: To maintain tissue viability and physiological function, the entire process of isolation and subsequent storage in Krebs solution at 4 °C should be completed within 6 h24,25,26,27. The solution should be replaced regularly during procedures to maintain oxygenation and tissue viability, while facilitating the clearance of blood components out of the vascular lumen.
  3. Visually identify the coronal plane by locating the renal hilum and aligning the cut to pass through both the renal pelvis and the lateral convex border. Using a sterile scalpel, bisect the kidney along the coronal plane to produce two symmetrical halves. This approach exposes the internal architecture, including the renal pyramids and columns, facilitating accurate identification and subsequent dissection of intrarenal arteries (Figure 1A).
    NOTE: In order to facilitate the separation of renal arteries, a portion of the renal pelvis, renal pyramids, and fat on the cross-sectional surface should be removed first to expose more blood vessels (Figure 1B). Be careful not to cut the blood vessels.
  4. Under a stereomicroscope, meticulously separate the interlobar, arcuate, and interlobular arteries using micro-dissection scissors and forceps in a 10 cm black-bottomed culture dish (Figure 1C).
    NOTE: In the human kidney, the interlobular artery courses through the renal pyramids, featuring the thickest vascular wall among the three arterial branches1,8, with an average diameter of approximately 1-2 mm. At the corticomedullary junction, the interlobar artery transitions into the arcuate artery, forming a semi-circular arc along the corticomedullary border, with an average diameter of approximately 500 µm. The blood vessels further branch into the renal cortex, which is the interlobular artery, with the thinnest vascular wall and smallest diameter among the three, about 200 µm (Figure 1C). Furthermore, when differentiating the three vascular branches, anatomical localization provides more critical information than absolute vascular diameter measurements1,8.
    1. Gently remove any surrounding tissue and fat from the arteries, making sure to clean them thoroughly in a 10 cm black-bottomed culture dish (Figure 1B).
  5. Use the isolated renal arteries (interlobular, arcuate, and interlobar arteries) for the following experiments:
    1. Prepare arterial segments for histological processing by carefully trimming isolated tissues to a standardized length of approximately 3 mm, which is suitable for paraffin embedding and histological sectioning. Fix each sample in 2 mL of 4% paraformaldehyde (PFA) at room temperature for 1 h to preserve tissue morphology. After fixation, process with standard dehydration, cleaning, and paraffin embedding protocols for subsequent hematoxylin and eosin (H&E) staining according to established histological protocols (Figure 2).
      NOTE: For potential histological assessments, the samples can undergo Masson's trichrome staining to evaluate collagen deposition or Verhoeff-Van Gieson (EVG) staining for elastic fiber visualization, according to established histological protocols28,29.
    2. For vascular function studies, store intrarenal rings in ice-cold Krebs solution until testing. The arterial rings should be completely submerged.
      NOTE: For subsequent molecular assays like western blot/qPCR, immediately snap-freeze fresh specimens in liquid nitrogen after dissection, then store at -80 °C following previously published procedures to preserve RNA integrity and prevent protein degradation30,31.

4. Arterial ring mounting

  1. Identify the interlobar, arcuate, and interlobular artery by their distinct anatomical positions (Figure 1C) and diameter ranges (Figure 2), then carefully section them into approximately 2-mm-long rings using microdissection scissors for vascular function studies (Figure 3A).
    NOTE: Immediately store each isolated arterial ring separately in 5 mL of ice-cold Krebs solution until functional testing. For functional testing, select unbranched vascular rings between 1.5-2.0 mm in length with clean perpendicular cuts. Exclude rings exceeding 2.0 mm to prevent mounting difficulties or rings under 1.5 mm to avoid unstable tension recordings.
  2. Prepare all arterial rings (from interlobar, arcuate, and interlobular arteries separately) as mentioned in step 4.1 and bring both Krebs and 60K+ solutions to room temperature. Then turn on the connected computer, followed by the wire myograph system, ensuring all components are properly initialized before proceeding with experiments.
  3. Before mounting the arterial rings in the wire myograph chambers, rinse each chamber twice with 5 mL of Krebs solution to ensure cleanliness and eliminate residual substances, then add 5 mL of Krebs solution to each chamber.
    1. Heat the solution to 37 °C using the myograph's built-in temperature control system. Simultaneously, continuously aerate the solution with gas (a mixture of 95% O2 and 5% CO2) via a connected gas line to maintain appropriate oxygenation and pH balance during the experiment.
  4. Prepare two 3 cm long guide wires with a diameter of 40 µm for each arterial ring and carefully transfer the isolated arterial rings to another 10 cm black-bottomed culture dish containing Krebs solution.
    NOTE: Different diameters of guide wires should be selected for blood vessels of different diameters. For the arterial ring with a diameter of about 100 µm or less, a 15 µm guide wire is required; if the diameter is between 100 µm and 200 µm, a 25 µm guide wire is used; if the diameter is greater than 200 µm, a 40 µm guide wire can be used. For the consistency of the experiment, it is recommended to use the same diameter for all vessels of the same type within a single experiment.19.
  5. Carefully insert the first guide wire into the arterial ring inside the dish, bend one side of the guide wire 90°, and then transfer it into the chamber for fixation on the clamp-type sample holder using the instrument-provided specialized screws for clockwise screw tightening (Figure 3B).
    NOTE: The guide wire is inserted to provide structural support during the transfer of the arterial ring when mounting it onto the sample holder, minimizing handling-induced damage or distortion.
  6. Before fixing the arterial ring on the holder, record the vessel length and width. Place the arterial ring between the two holders (Figure 3C) and read the micrometer scale, where 1 scale represents 10 µm. Subtract the scale value measured when the holders barely touch each other (Figure 3D) to determine the arterial ring's length and width.
  7. Thread another guide wire through the blood vessel and finally fix it on the other side. Wind the guide wire clockwise around the fixing screws on both sides and ensure it is tightly attached to the sample holder surface.
    NOTE: During the entire fixation process, it is necessary to avoid pulling the blood vessel or scratching the inner wall of the blood vessel with the guide wire to avoid damaging the vascular endothelium. The guide wire should be wrapped around the fixing screw in a single layer without overlapping. The two guide wires should be on the same horizontal plane, parallel to each other, and not intersect with each other. In addition, after fixing the guide wire, adjust the sample holder on both sides closer; however, they should be close to each other without actually contacting, so as to avoid force, as shown in Figure 3E and Figure 3F.
  8. Install the chamber back into the wire myograph system. Then, cover the lid and ventilate the gas. Connect the sensor and open the data acquisition software for vascular tension testing.

5. Vessel-specific normalization for optimal initial tension determination

NOTE: Establishing stable baseline conditions, defined as the optimal resting tension for maximal contractile responsiveness, is critical for reproducible functional measurements. However, due to inherent inter-individual physiological differences, human blood vessels exhibit greater variability in baseline mechanical properties compared to animal blood vessels32,33,34,35,36. These factors, combined with intrinsic vascular heterogeneity (e.g., structural and functional disparities in diameter, wall thickness, and contractile properties), collectively lead to significant variations in baseline tension. Therefore, to ensure experimental reproducibility, it is recommended to normalize each blood vessel individually. The wire myograph system includes a dedicated normalization module, which enables precise determination of the baseline tension for each vessel.

  1. Mount the arterial rings in the chamber as described in steps 4.1 to 4.8. Begin chart recording, zero the channel (Figure 4A), allow stabilization for at least 30 min, then re-zero the channel again before proceeding with normalization.
  2. Select Normalization Settings from DMT menu (Figure 4B) and set up the parameters as follows (Figure 4C):
    1. Eyepiece calibration (mm/div): 1; Target pressure (kPa): 13.3; IC1/IC100: 0.9; Online averaging time (seconds): 3; Delay time (seconds): 60.
      NOTE: IC1 denotes the internal vessel circumference at maximal active force production, while IC100 represents the circumference corresponding to 100 mmHg transmural pressure. Human renal arteries exhibit substantial inter-specimen variability, requiring standardization, unlike animal models that have consistent vascular properties for experimental determination and reuse of IC1/IC100 ratios. Due to significant variations in vessel diameter, wall thickness, and mechanical properties between individual human specimens, each vessel must undergo independent normalization to establish its unique baseline characteristics. The ratio of 0.9 was therefore established as a placeholder for protocol initialization here and not used in practice in this protocol.
  3. Select the channel corresponding to the target artery and open the normalization screen from the DMT menu (Figure 4C), populate it with the appropriate data:
    1. Tissue end points: a1 should be 0, a2 should be the length of the vessel (in mm); wire diameter: 40 µm (input the diameter of the wire used); micrometer reading: the value from the micrometer scale, click Add point to record the point.
  4. When ready, click Add point to record the first point and start the normalization procedure.
  5. Apply passive stretch, wait for 3 min, enter the micrometer reading as the next point, then add 5 mL of 60K+ to elicit a potassium-mediated vessel contraction (Figure 5).
  6. Wait until the contraction reaches a plateau. Then, wash out the 60K+ solution with 5 mL of Krebs solution 3 times, and calculate the active force data from the trace by subtracting the passive force at each stretch from the potassium-activated force (Figure 5).
  7. Repeat steps 5.5 to 5.6 ("Stretch-60K+ stimulation-Wash" steps) until the active force reaches its maximum, then determine the optimal baseline tension at which the vessel's active force production is maximal (Figure 5).
  8. Equilibrate the arterial ring in the chamber for 10 min before proceeding to the next steps of the experiment.

6. Detection of renal artery ring reactivity

  1. Make sure the volume of Krebs in each chamber is 5 mL before the start of the experiment.
  2. To evaluate phenylephrine-induced concentration-dependent contraction, apply cumulative Phe in half-log increments (10-9 to 10-4 M)37.
  3. Start with the lowest Phe concentration (10-9 M). Add 5 µL of Phe stock (10-6 M) to the chamber containing 5 mL of Krebs buffer, continuously monitor and wait until contraction stabilizes (2-3 min of unchanged tension), mark the point, then add the next concentration in a half-log increment. Repeat until reaching the final concentration (10-4 M), maintaining consistent monitoring throughout (Figure 6).
  4. For additional drug testing, wash the chamber with warm 5 mL of Krebs solution at least 5 times until tension returns to baseline and remains stable for at least 10 min.
  5. To evaluate acetylcholine-induced concentration-dependent vasodilation, apply cumulative ACh in half-log increments (10-8 to 10-5.5 M).
    NOTE: In normal vasculature, the response to acetylcholine (ACh) serves as a functional indicator of endothelial integrity37,38. The vasodilatory response to ACh reflects endothelial function through NO-mediated relaxation in healthy vessels. However, under pathological conditions, this response may be attenuated or abolished due to impaired nitric oxide (NO) bioavailability and/or altered receptor signaling pathways.
  6. Pre-contract vessels with 10-4 M Phe, add 5 µL of Phe stock (10-1 M) to the chamber containing 5 mL of Krebs buffer, and incubate for 15 min. Starting from the lowest ACh concentration, add 5 µL of Ach stock (10-5 M) to the chamber containing 5 mL of Krebs buffer, continuously monitor and wait for vasodilation response (confirmed by sustained trace detection), mark the point, then proceed to the next concentration. Repeat until all are tested (Figure 7).
  7. Complete the experiment by saving the file and removing the arterial rings. Thoroughly clean the chamber by incubating with 10 mL of 8% acetic acid solution for 3 min, then rinse with 10 mL of water 3 times. Next, incubate in 10 mL of 95% ethanol for 3 min and rinse again with 10 mL of water 3 times. Finally, turn off both the heater and gas, ensuring all liquid is removed before shutting off the gas.

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Results

In the human kidney, the interlobar artery runs between renal pyramids alongside its corresponding vein. Characterized by a relatively thick vascular wall and significant adipose tissue encasement, this artery requires particularly careful dissection. The surrounding adipose tissue should be removed meticulously to avoid application of excessive force during the isolation procedure (Figure 1C).

The arcuate artery is anatomically positioned at the corticomedullary ...

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Discussion

Renal perfusion stability serves as both a critical therapeutic target and a window into hemodynamic pathophysiology39,40. We present a protocol for hierarchical isolation and functional assessment of human renal arteries from fresh nephrectomy specimens, addressing significant gaps in current methodologies. Unlike existing approaches limited to cadaveric specimens or animal models, our technique enables reliable isolation of viable intrarenal arteries (interloba...

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Disclosures

The authors have nothing to disclose.

Acknowledgements

The authors would like to thank the volunteers for their participation in this study, and also thank the surgical staff at Urology Department of Peking University First Hospital for providing kidney tissue. This work was supported by the Beijing Municipal Natural Science Foundation (F251013 to Y. Z., 7232096 to Y. L.); National Science Foundation of China (82325004, 92168114 to Y. Z., 82170422 to Y. L.); National Key R&D Program of China (2021YFF0501401, 2018YFA0800501 to Y. Z., 2021YFF0501404 to Y. L., 2023YFC2415500 to L. Y.); Research Project of Peking University Third Hospital in State Key Laboratory of Vascular Homeostasis and Remodeling (Peking University; 2024-VHR-SY-07 to Y.Z.); 2023 Beijing Municipal Health Commission Capital Medical Science and Technology Innovation Achievement Transformation Excellent Promotion Program Project (YC202301QX0162 to L. Y.); National High Level Hospital Clinical Research Funding (Scientific and Technological Achievements Transformation Incubation Guidance Fund Project of Peking University First Hospital, 2024CX24 to L. Y.).

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
Acetylcholine chlorideSigma-Aldrich, Merck, Darmstadt, GermanyA6625
Black bottomed culture dishDanish Myo Technology, Aarhus, Denmark300412
CaCl2Sangon Biotech Co.,Ltd.,Shanghai, ChinaA501330
D-glucoseSangon Biotech Co.,Ltd.,Shanghai, ChinaA610219
Dumont forcepsDanish Myo Technology, Aarhus, Denmark300413
Geuder dissection scissorDanish Myo Technology, Aarhus, Denmark400431
Guide wireDanish Myo Technology, Aarhus, Denmark400447Diameter 40 µm
KClSangon Biotech Co.,Ltd.,Shanghai, ChinaA100395
KH2PO4Sangon Biotech Co.,Ltd.,Shanghai, ChinaA100781
LabChart Professional version 8.3 ADInstruments, Australia-
MgCl2·6H2OSangon Biotech Co.,Ltd.,Shanghai, ChinaA100288
Multi myograph system Danish Myo Technology, Aarhus, Denmark620M
NaClSangon Biotech Co.,Ltd.,Shanghai, ChinaA100241
NaHCO3Sangon Biotech Co.,Ltd.,Shanghai, ChinaA100865
PhenylephrineSigma-Aldrich, Merck, Darmstadt, GermanyP6126

References

  1. Molema, G., Aird, W. C. Vascular heterogeneity in the kidney. Semin Nephrol. 32 (2), 145-155 (2012).
  2. Jentzer, J. C., et al. Contemporary management of severe acute kidney injury and refractory cardiorenal syndrome: Jacc council perspectives. J Am Coll Cardiol. 76 (9), 1084-1101 (2020).
  3. Kanji, H. D., et al. Difference between preoperative and cardiopulmonary bypass mean arterial pressure is independently associated with early cardiac surgeryassociated acute kidney injury. J Cardiothorac Surg. 5 (1), 71(2010).
  4. Kwiatkowska, E., Kwiatkowski, S., Dziedziejko, V., Tomasiewicz, I., Domanski, L. Renal microcirculation injury as the main cause of ischemic acute kidney injury development. Biology (Basel). 12 (2), 327(2023).
  5. Ruilope, L. M. Hypertension in 2010:Blood pressure and the kidney. Nat Rev Nephrol. 7 (2), 73-74 (2011).
  6. Barger, A. C., Herd, J. A. The renal circulation. N Engl J Med. 284 (9), 482-490 (1971).
  7. Beloncle, F., Piquilloud, L., Asfar, P. Critical care nephrology. , third edition, Elsevier. Philadelphia. (2019).
  8. MompeóCorredera, B., HernándezMorera, P., CastañoGonzález, I., QuintanaMontesdeoca, M. D. P., MederosReal, N. Regions of the human renal artery:Histomorphometric analysis. Anat Cell Biol. 55 (3), 330-340 (2022).
  9. Evans, R. G., Smith, D. W., Lee, C. J., Ngo, J. P., Gardiner, B. S. What makes the kidney susceptible to hypoxia. Anat Rec (Hoboken). 303 (10), 2544-2552 (2020).
  10. Hwa, J. J., Ghibaudi, L., Williams, P., Chatterjee, M. Comparison of acetylcholinedependent relaxation in large and small arteries of rat mesenteric vascular bed. Am J Physiol. 266 (3 Pt 2), H952-H958 (1994).
  11. Buus, N. H., Van Bavel, E., Mulvany, M. J. Differences in sensitivity of rat mesenteric small arteries to agonists when studied as ring preparations or as cannulated preparations. Br J Pharmacol. 112 (2), 579-587 (1994).
  12. Barone, R., et al. Renal arterial and venous doppler in cardiorenal syndrome: Pathophysiological and clinical insights. Biomedicines. 12 (6), 1166(2024).
  13. Cuttone, G., et al. Exploring the utility of renal resistive index in critical care: Insights into ARDS and cardiac failure. Biomedicines. 13 (2), 519(2025).
  14. Li, Y., et al. Clinical practice report of contrastenhanced ultrasound in renal artery disease. J Ultrasound Med. 43 (1), 117-125 (2023).
  15. Andersen, F. T., et al. Continuous assessment of tissue perfusion using quantitative indocyanine green fluorescence imaging during controlled hypo and reperfusion. Surg Endosc. 39 (10), 6512-6521 (2025).
  16. Schubert, R., Gaynullina, D., Shvetsova, A., Tarasova, O. S. Myography of isolated blood vessels:Considerations for experimental design and combination with supplementary techniques. Front Physiol. 14 (8), 1176748(2023).
  17. Wang, Z., et al. Mir1865p carried by M2 macrophagederived exosomes downregulates TRPP2 expression in airway smooth muscle to alleviate asthma progression. Int Immunopharmacol. 148 (1), 114107(2025).
  18. Ikeda, M., Nakada, A., Abukawa, H., Yamazaki, T., Maruyama, I. Vibegron inhibits enhanced spontaneous contractions induced by anoxia/reoxygenation in isolated whole bladder from rats. Eur J Pharmacol. 926 (1), 175017(2022).
  19. Wenceslau, C. F., et al. Guidelines for the measurement of vascular function and structure in isolated arteries and veins. Am J Physiol Heart Circ Physiol. 321 (1), H77-H111 (2021).
  20. Bayliss, W. M. On the local reactions of the arterial wall to changes of internal pressure. J Physiol. 28 (3), 220-231 (1902).
  21. Darmon, M., Schnell, D., Schneider, A. Monitoring of renal perfusion. Intensive Care Med. 48 (10), 1505-1507 (2022).
  22. Offenhäuser, C., et al. EphA2 regulates vascular permeability and prostate cancer metastasis via modulation of cell junction protein phosphorylation. Oncogene. 44 (4), 208-227 (2025).
  23. Vitverova, B., et al. Soluble endoglin and hypercholesterolemia aggravate endothelial and vessel wall dysfunction in mouse aorta. Atherosclerosis. 271, 15-25 (2018).
  24. Maathuis, M. H. J., De Groot, M., Ploeg, R. J., Leuvenink, H. G. D. Deterioration of endothelial and smooth muscle cell function in DCD kidneys after static cold storage in IGL1 or UW. J Surg Res. 152 (2), 231-237 (2009).
  25. Ingemansson, R., Budrikis, A., Bolys, R., Sjoberg, T., Steen, S. Effect of temperature in longterm preservation of vascular endothelial and smooth muscle function. Ann Thorac Surg. 61 (5), 1413-1417 (1996).
  26. McIntyre, C. A., Williams, B. C., Lindsay, R. M., McKnight, J. A., Hadoke, P. W. Preservation of vascular function in rat mesenteric resistance arteries following cold storage, studied by small vessel myography. Br J Pharmacol. 123 (8), 1555-1560 (1998).
  27. Ingemansson, R., Sjoberg, T., Massa, G., Steen, S. Longterm preservation of vascular endothelium and smooth muscle. Ann Thorac Surg. 59 (5), 1177-1181 (1995).
  28. Zheng, Y., et al. HMGB2 promotes smooth muscle cell proliferation through PPARγ/PGC1α pathwaymediated glucose changes in aortic dissection. Atherosclerosis. 399, 119044(2024).
  29. Zhang, B., Zhang, Y., Deng, F., Fang, S. Ligustrazine prevents basilar artery remodeling in twokidneytwoclip renovascular hypertension rats via suppressing PI3K/AKT signaling. Microvasc Res. 128, 103938(2020).
  30. Somiari, S. B., et al. Assessing the quality of RNA isolated from human breast tissue after ambient room temperature exposure. PLoS One. 17 (1), e0262654(2022).
  31. Srinivasan, M., Sedmak, D., Jewell, S. Effect of fixatives and tissue processing on the content and integrity of nucleic acids. Am J Pathol. 161 (6), 1961-1971 (2002).
  32. Potente, M., Makinen, T. Vascular heterogeneity and specialization in development and disease. Nat Rev Mol Cell Biol. 18 (8), 477-494 (2017).
  33. Lake, B. B., et al. An atlas of healthy and injured cell states and niches in the human kidney. Nature. 619 (7970), 585-594 (2023).
  34. Greenbaum, S., et al. A spatially resolved timeline of the human maternal-fetal interface. Nature. 619 (7970), 595-605 (2023).
  35. The Tabula Sapiens Consortium. The Tabula Sapiens: A multipleorgan, singlecell transcriptomic atlas of humans. Science. 376 (6594), eabl4896(2022).
  36. Barnett, S. N., et al. An organotypic atlas of human vascular cells. Nat Med. 30 (12), 3468-3481 (2024).
  37. Shen, Y., et al. Targeting cytokinelike protein FAM3D lowers blood pressure in hypertension. Cell Rep Med. 4 (6), 101072(2023).
  38. Zhao, L., et al. Restoration of autophagic flux improves endothelial function in diabetes through lowering mitochondrial ROSmediated eNOS monomerization. Diabetes. 71 (5), 1099-1114 (2022).
  39. Ergin, B., Akin, S., Ince, C. Kidney microcirculation as a target for innovative therapies in AKI. J Clin Med. 10 (18), (2021).
  40. Deferrari, G., Cipriani, A., La Porta, E. Renal dysfunction in cardiovascular diseases and its consequences. J Nephrol. 34 (1), 137-153 (2021).
  41. Owen, M. K., et al. Perivascular adipose tissue potentiates contraction of coronary vascular smooth muscle: Influence of obesity. Circulation. 128 (1), 9-18 (2013).
  42. AyalaLopez, N., et al. Perivascular adipose tissue contains functional catecholamines. Pharmacol Res Perspect. 2 (3), e00041(2014).
  43. AyalaLopez, N., Thompson, J. M., Watts, S. W. Perivascular adipose tissue's impact on norepinephrineinduced contraction of mesenteric resistance arteries. Front Physiol. 8, 37(2017).
  44. Ahmad, M. F., et al. Perivascular adipocytes store norepinephrine by vesicular transport. Arterioscler Thromb Vasc Biol. 39 (2), 188-199 (2019).
  45. Ya-Jing, W., et al. Adipose tissue-derived cytokines, CTRPs as biomarkers and therapeutic targets in metabolism and the cardiovascular system. Vessel Plus. (1), 202-212 (2017).
  46. Osol, G., Brekke, J. F., McElroyYaggy, K., Gokina, N. I. Myogenic tone, reactivity, and forced dilatation: A threephase model of in vitro arterial myogenic behavior. Am J Physiol Heart Circ Physiol. 283 (6), H2260-H2267 (2002).

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Microdissection TechniqueWire MyographyVascular DysfunctionIntrarenal Artery IsolationVasomotor ResponsesMuscle Length-TensionHematoxylin Eosin StainingPhenylephrine ContractionArterial Normalization
Multilevel Microdissection and Functional-Structural Profiling of Human Renal Arterial Branches
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