The authors thank the Danish Molecular Biomedical Imaging Center at the Faculty of Natural Sciences, University of Southern Denmark, for the use of laboratories and microscopes. Kristoffer Rosenstand and Ulla Melchior are acknowledged for excellent technical assistance with the pressure myography and imaging.

The authors have nothing to disclose.

This work represents our suggestion for a standardized, combined imaging and pressure myography approach, valuable for simultaneous assessment of the mechanical properties of resistance arteries and pressure-related changes in the structure of the arterial wall over a pressure range from 0 to 100 mmHg. The presented approach was developed using custom built equipment, however, any pressure myograph that fits on a two-photon excitation fluorescence microscope can be used, when the design of both equipments allows imaging ...

The pathogenic contribution and effects of resistance artery remodeling are documented in essential hypertension, diabetes and the metabolic syndrome1,2,3,4,5. Deciphering the relationship between the mechanical and microarchitectural properties of the microvascular wall is essential for developing mathematical models of this association. Such models will improve understanding the remodeling process and will support the development of in silico models useful for testing pharmacological strategies targeting disease related remodeling of the arterial wall.
Prior studies focused in understanding how the microarchitecture of the arterial wall relates to arterial wall mechanics by incorporating mechanical measures and the microarchitecture of the extracellular matrix (ECM) are almost exclusively performed on large, elastic conduit arteries from mice or swine6,7,8,9,10,11. Imaging of the microstructures of the wall is typically performed using nonlinear optical techniques, taking advantage of the autofluorescence of elastin and second harmonic generation by collagen. This allows spatiotemporal imaging of the two major components of the extracellular matrix, elastin and collagen, without a need for staining. Imaging of the arterial wall in full thickness is a challenge in large conduit arteries due to scatter of the light in the thick tunica media. However, to determine how the microarchitecture of the structural components of the arterial wall relate to the observed mechanical properties, three-dimensional information must be obtained during the mechanical testing. For large arteries like the human aorta, this requires biaxial mounting, mechanical testing and imaging of regions of interest in 1-2 cm2 pieces of the arterial wall7,9,10,12. Only part of the wall can be imaged and mechanically tested.
For smaller arteries of any species (e.g., human pericardial13, pulmonary14 and subcutaneous15 arteries, rat mesenteric arteries16,17,18,19,20, mouse cremaster, mesenteric, cerebral, femoral and carotid arteries21,22,23,24,25,26,27) imaging of the entire wall thickness is possible and can be combined with mechanical testing. This allows simultaneous recording of the mechanical properties and the structural arrangements within the wall. However, a direct mathematical modeling of the relationship between the observed alterations in the three-dimensional structure of the ECM and changed mechanical properties of the resistance arterial wall, has to the best of our knowledge only been reported upon recently in human resistance arteries13,15.
In this work, an ex vivo method for passive mechanical testing and simultaneous three-dimensional imaging of the microarchitecture of elastin and collagen in the arterial wall of isolated human resistance arteries is described. The imaging protocol can be applied to resistance arteries of any species of interest. Image analyses are described for obtaining measures of internal elastic lamina branching angles and adventitial collagen straightness13 using Fiji28. Collagen and elastin volume densities are determined using Ilastik software29 and finally, the inclusion of the mechanical and imaging data in mathematical models of the arterial wall mechanics is discussed.
The goal of describing the imaging and image analyses techniques in combination with mathematical modeling is to provide investigators a systematical approach to describe and understand observed pressure induced changes in the ECM of resistance arteries. The described method is focused in quantifying the changes in the ECM in a vessel during pressurization, by comparing the structure of the ECM at 20, 40 and 100 mmHg. These pressures were chosen for determining the structure of the arterial wall at its more compliant (20 mmHg), stiff (100 mmHg) and intermediate (40 mmHg) state, respectively. However, any process in the vascular wall of live arteries, including changes induced by vasoactive components, hysteresis and flow, can be quantified, depending on the research hypothesis in question by the investigator.
The use of two-photon excitation fluorescence microscopy (TPEM) in combination with a pressure myograph for studying pressure (or other) induced changes in the ECM of live arteries is emphasized. First, because this allows simultaneous acquisition of the overall three-dimensional structure of the arterial wall (diameter and wall thickness) along with three-dimensional label-free acquisition of high quality, detailed images of the collagen and elastin microarchitectures as described13 by taking advantage of the elastin autofluorescence and the collagen second harmonic generation signal (SHG)30. Second, TPEM allows use of low-energy near-infrared excitation light, minimizing photodamage of the tissue and thus, repeated imaging at exactly the same position within the vascular wall is allowed, permitting repeated-measurements analyses of observed changes.
The use of an alternative approach using confocal imaging of pressure fixed arteries is discussed to allow users without access to TPEM an opportunity to use the described method as well. Information on ECM structure and volume densities can also be retrieved from two-dimensional analyses of tissues sectioned in serial, e.g. as described by31,32. However, due to the lack of possibility to retrieve three-dimensional structural information over the length scales of the artery as well as during changing conditions using this method, it is not recommend using this approach for investigations of pressure and treatment induced three-dimensional changes in the ECM.
The minimum requirement for the investigator to apply the herein described method is the access to a setup for cannulation and pressurization of arteries in combination with a confocal or two-photon excitation fluorescence microscope. The setup described in the following protocol is a custom-built pressure myograph with a longitudinal force transducer, built to fit on a custom built inverted two-photon excitation fluorescence microscope.

<table><tbody><tr><td>Fine Science Tools</td><td>15401-12</td><td></td><td></td></tr><tr><td>Fine Science Tools</td><td>11251-23</td><td></td><td></td></tr><tr><td>Nikon</td><td>SMZ800N</td><td></td><td></td></tr><tr><td>&amp;nbsp;Sigma-Aldrich, Br&amp;oslash;ndby, Denmark.</td><td>761028</td><td>for dissection purpose</td><td></td></tr><tr><td>Vitrex Medical A/S, Herlev, Denmark</td><td>1.63, 2.13, 210mm</td><td></td><td></td></tr><tr><td>Smiths medical Intl, UK</td><td></td><td></td><td></td></tr><tr><td>Ethicon</td><td>Ethilon 11-0</td><td></td><td></td></tr><tr><td>Custom built</td><td></td><td>DK patent number 201200167, University of Southern Denmark, J. Schoubo V. Jensen, F. Jensen. T.R. Uhrenholt</td><td></td></tr><tr><td>Mettler toledo</td><td></td><td></td><td></td></tr><tr><td>&amp;nbsp;Sigma-Aldrich, Br&amp;oslash;ndby, Denmark.</td><td>B3259</td><td></td><td></td></tr><tr><td>&amp;nbsp;Sigma-Aldrich, Br&amp;oslash;ndby, Denmark.</td><td>A7030</td><td></td><td></td></tr><tr><td>&amp;nbsp;Sigma-Aldrich, Br&amp;oslash;ndby, Denmark.</td><td>C5670</td><td></td><td></td></tr><tr><td>&amp;nbsp;Sigma-Aldrich, Br&amp;oslash;ndby, Denmark.</td><td>G7021</td><td></td><td></td></tr><tr><td>&amp;nbsp;Sigma-Aldrich, Br&amp;oslash;ndby, Denmark.</td><td>E3889</td><td></td><td></td></tr><tr><td>Merck Millipore, Hellerup, Denmark</td><td>1.00496.9010</td><td>Phosphate buffered (pH 6.9) 4% formaldehyde solution&amp;nbsp;</td><td></td></tr><tr><td>&amp;nbsp;Sigma-Aldrich, Br&amp;oslash;ndby, Denmark.</td><td>H3784</td><td></td><td></td></tr><tr><td>&amp;nbsp;Sigma-Aldrich, Br&amp;oslash;ndby, Denmark.</td><td>P9666</td><td></td><td></td></tr><tr><td>&amp;nbsp;Sigma-Aldrich, Br&amp;oslash;ndby, Denmark.</td><td>P5655</td><td></td><td></td></tr><tr><td>&amp;nbsp;Sigma-Aldrich, Br&amp;oslash;ndby, Denmark.</td><td>M2643</td><td></td><td></td></tr><tr><td>&amp;nbsp;Sigma-Aldrich, Br&amp;oslash;ndby, Denmark.</td><td>S2002</td><td></td><td></td></tr><tr><td>&amp;nbsp;Sigma-Aldrich, Br&amp;oslash;ndby, Denmark.</td><td>S5886</td><td></td><td></td></tr><tr><td>&amp;nbsp;Sigma-Aldrich, Br&amp;oslash;ndby, Denmark.</td><td>S5761</td><td></td><td></td></tr><tr><td>&amp;nbsp;Sigma-Aldrich, Br&amp;oslash;ndby, Denmark.</td><td>1.06462</td><td></td><td></td></tr><tr><td>Gibco, ThermoFisher Scientific</td><td>10010015</td><td></td><td></td></tr><tr><td>&amp;nbsp;Sigma-Aldrich, Br&amp;oslash;ndby, Denmark.</td><td>PHR1423</td><td></td><td></td></tr><tr><td>&amp;nbsp;Sigma-Aldrich, Br&amp;oslash;ndby, Denmark.</td><td>Z370525</td><td></td><td></td></tr><tr><td>&amp;nbsp;Tocris Bioscience, Bristol, UK</td><td>538944</td><td></td><td></td></tr><tr><td>Nikon</td><td>Custom built</td><td></td><td></td></tr><tr><td>Spectra Physics, Mountain View, CA</td><td></td><td></td><td></td></tr><tr><td>Nikon</td><td>CFI Plan Apo IR SR 60XWI NA 1.27</td><td></td><td></td></tr><tr><td>Nikon</td><td>CFI Plan Fluor 20XMI (multi-immersion) NA 0.75</td><td></td><td></td></tr><tr><td>Hamamatsu, Ballerup, Denmark</td><td>H7422P-40</td><td></td><td></td></tr><tr><td>AHF analysentechnik AG (T&amp;uuml;bingen, Germany).</td><td>ChromaET 460 nm long pass dichroic</td><td></td><td></td></tr><tr><td>AHF analysentechnik AG (T&amp;uuml;bingen, Germany).</td><td>Semrock FF01-520/35-25 BrightLine filter</td><td></td><td></td></tr><tr><td>AHF analysentechnik AG (T&amp;uuml;bingen, Germany).</td><td>Chroma ET402/15X&amp;nbsp;</td><td></td><td></td></tr><tr><td>Scotch TM</td><td></td><td></td><td></td></tr><tr><td></td><td>coverslip thickness should match used objective on microscope (#1 or #1.5), alternatively, set adjustment collar to match coverslip</td><td></td><td></td></tr></tbody></table>

assessing collagen and elastin pressure-dependent microarchitectures in live, human resistance arteries by label-free fluorescence microscopy

The pathogenic contribution of resistance artery remodeling is documented in essential hypertension, diabetes and the metabolic syndrome. Investigations and development of microstructurally motivated mathematical models for understanding the mechanical properties of human resistance arteries in health and disease have the potential to aid understanding how disease and medical treatments affect the human microcirculation. To develop these mathematical models, it is essential to decipher the relationship between the mechanical and microarchitectural properties of the microvascular wall. In this work, we describe an ex vivo method for passive mechanical testing and simultaneous label-free three-dimensional imaging of the microarchitecture of elastin and collagen in the arterial wall of isolated human resistance arteries. The imaging protocol can be applied to resistance arteries of any species of interest. Image analyses are described for quantifying i) pressure-induced changes in internal elastic lamina branching angles and adventitial collagen straightness using Fiji and ii) collagen and elastin volume densities determined using Ilastik software. Preferably all mechanical and imaging measurements are performed on live, perfused arteries, however, an alternative approach using standard video-microscopy pressure myography in combination with post-fixation imaging of re-pressurized vessels is discussed. This alternative method provides users with different options for analysis approaches. The inclusion of the mechanical and imaging data in mathematical models of the arterial wall mechanics is discussed, and future development and additions to the protocol are proposed.

The custom-built pressure myograph for imaging used in this work is shown in Figure 1. Special attention for the design of the myograph was paid to i) the chamber with a small volume (2 mL) and ii) the possibility for positioning the cannulae close to, and parallel with the glass bottom (Figure 1B). The bottom of the chamber fits a 50 &#215; 24 mm #1.5 glass coverslip (replaceable). The pressure controller was built from a standa...

Watch this Scientific Journal Video about Assessing Collagen and Elastin Pressure-dependent Microarchitectures in Live, Human Resistance Arteries by Label-free Fluorescence Microscopy at JoVE.com

Assessing Collagen and Elastin Pressure-dependent Microarchitectures in Live, Human Resistance Arteries by Label-free Fluorescence Microscopy

We describe simultaneous mechanical testing and 3D-imaging of the arterial wall of isolated, live human resistance arteries, and Fiji and Ilastik image analyses for the quantification of elastin and collagen spatial organization and volume densities. We discuss the use of these data in mathematical models of arterial wall mechanics.

The pathogenic contribution of resistance artery remodeling is documented in essential hypertension, diabetes and the metabolic syndrome. Investigations ...

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Cell Biology

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Unit 1

Extracellular Matrix in Animals

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10.5K Views.  University of Southern Denmark. We beschrijven gelijktijdige mechanische testen en 3D-imaging van de arteriële muur van geïsoleerd levende menselijke weerstand slagaders, en Fiji en Ilastik analyses van de afbeelding voor de kwantificering van elastine en collageen ruimtelijke organisatie en volume dichtheden. We bespreken het gebruik van deze gegevens in wiskundige modellen van arteriële muur mechanica.

Video: Assessing Collagen and Elastin Pressure-dependent Microarchitectures in Live, Human Resistance Arteries by Label-free Fluorescence Microscopy

Live-cell Imaging of Migrating Cells Expressing Fluorescently-tagged Proteins in a Three-dimensional Matrix

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound healing, tissue regeneration, and cancer metastasis, cells must navigate through complex, three-dimensional extracellular tissue. To better understand the mechanisms behind these biological processes, it is important to examine the roles of the proteins responsible for driving cell migration. Here, we outline a protocol to study the mechanisms of cell migration using the epithelial cell line (MDCK), and a three-dimensional, fibrous, self-polymerizing matrix as a model system. This optically clear extracellular matrix is easily amenable to live-cell imaging studies and better mimics the physiological, soft tissue environment. This report demonstrates a technique for directly visualizing protein localization and dynamics, and deformation of the surrounding three-dimensional matrix. 
Examination of protein localization and dynamics during cellular processes provides key insight into protein functions. Genetically encoded fluorescent tags provide a unique method for observing protein localization and dynamics. Using this technique, we can analyze the subcellular accumulation of key, force-generating cytoskeletal components in real-time as the cell maneuvers through the matrix. In addition, using multiple fluorescent tags with different wavelengths, we can examine the localization of multiple proteins simultaneously, thus allowing us to test, for example, whether different proteins have similar or divergent roles. Furthermore, the dynamics of fluorescently tagged proteins can be quantified using Fluorescent Recovery After Photobleaching (FRAP) analysis. This measurement assays the protein mobility and how stably bound the proteins are to the cytoskeletal network.
By combining live-cell imaging with the treatment of protein function inhibitors, we can examine in real-time the changes in the distribution of proteins and morphology of migrating cells. Furthermore, we also combine live-cell imaging with the use of fluorescent tracer particles embedded within the matrix to visualize the matrix deformation during cell migration. Thus, we can visualize how a migrating cell distributes force-generating proteins, and where the traction forces are exerted to the surrounding matrix. Through these techniques, we can gain valuable insight into the roles of specific proteins and their contributions to the mechanisms of cell migration.

Cellular processes such as cell migration have traditionally been studied on two-dimensional, stiff plastic surfaces. This report describes a technique for directly visualizing protein localization and analyzing protein dynamics in cells migrating in a more physiologically relevant, three-dimensional matrix.

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound ...

Non-contact, Label-free Monitoring of Cells and Extracellular Matrix using Raman Spectroscopy

Non-destructive, non-contact and label-free technologies to monitor cell and tissue cultures are needed in the field of biomedical research.1-5 However, currently available routine methods require processing steps and alter sample integrity. Raman spectroscopy is a fast method that enables the measurement of biological samples without the need for further processing steps. This laser-based technology detects the inelastic scattering of monochromatic light.6 As every chemical vibration is assigned to a specific Raman band (wavenumber in cm-1), each biological sample features a typical spectral pattern due to their inherent biochemical composition.7-9 Within Raman spectra, the peak intensities correlate with the amount of the present molecular bonds.1 Similarities and differences of the spectral data sets can be detected by employing a multivariate analysis (e.g. principal component analysis (PCA)).10
Here, we perform Raman spectroscopy of living cells and native tissues. Cells are either seeded on glass bottom dishes or kept in suspension under normal cell culture conditions (37 &deg;C, 5% CO2) before measurement. Native tissues are dissected and stored in phosphate buffered saline (PBS) at 4 &deg;C prior measurements. Depending on our experimental set up, we then either focused on the cell nucleus or extracellular matrix (ECM) proteins such as elastin and collagen. For all studies, a minimum of 30 cells or 30 random points of interest within the ECM are measured. Data processing steps included background subtraction and normalization.

Raman spectroscopy is a suitable technique for the non-contact, label-free analysis of living cells, tissue-engineered constructs and native tissues. Source-specific spectral fingerprints can be generated and analyzed using multivariate analysis.

Non-destructive, non-contact and label-free technologies to monitor cell and tissue cultures are needed in the field of biomedical research.1-5 However, ...

Imaging Leukocyte Adhesion to the Vascular Endothelium at High Intraluminal Pressure

Worldwide, hypertension is reported to be in approximately a quarter of the population and is the leading biomedical risk factor for mortality worldwide. In the vasculature hypertension is associated with endothelial dysfunction and increased inflammation leading to atherosclerosis and various disease states such as chronic kidney disease2, stroke3 and heart failure4. An initial step in vascular inflammation leading to atherogenesis is the adhesion cascade which involves the rolling, tethering, adherence and subsequent transmigration of leukocytes through the endothelium. Recruitment and accumulation of leukocytes to the endothelium is mediated by an upregulation of adhesion molecules such as vascular cell adhesion molecule-1 (VCAM-1), intracellular cell adhesion molecule-1 (ICAM-1) and E-selectin as well as increases in cytokine and chemokine release and an upregulation of reactive oxygen species5. In vitro methods such as static adhesion assays help to determine mechanisms involved in cell-to-cell adhesion as well as the analysis of cell adhesion molecules. Methods employed in previous in vitro studies have demonstrated that acute increases in pressure on the endothelium can lead to monocyte adhesion, an upregulation of adhesion molecules and inflammatory markers6 however, similar to many in vitro assays, these findings have not been performed in real time under physiological flow conditions, nor with whole blood. Therefore, in vivo assays are increasingly utilised in animal models to demonstrate vascular inflammation and plaque development. Intravital microscopy is now widely used to assess leukocyte adhesion, rolling, migration and transmigration7-9. When combining the effects of pressure on leukocyte to endothelial adhesion the in vivo studies are less extensive. One such study examines the real time effects of flow and shear on arterial growth and remodelling but inflammatory markers were only assessed via immunohistochemistry10. Here we present a model for recording leukocyte adhesion in real time in intact pressurised blood vessels using whole blood perfusion. The methodology is a modification of an ex vivo vessel chamber perfusion model9 which enables real-time analysis of leukocyte -endothelial adhesive interactions in intact vessels. Our modification enables the manipulation of the intraluminal pressure up to 200 mmHg allowing for study not only under physiological flow conditions but also pressure conditions. While pressure myography systems have been previously demonstrated to observe vessel wall and lumen diameter11 as well as vessel contraction this is the first time demonstrating leukocyte-endothelial interactions in real time. Here we demonstrate the technique using carotid arteries harvested from rats and cannulated to a custom-made flow chamber coupled to a fluorescent microscope. The vessel chamber is equipped with a large bottom coverglass allowing a large diameter objective lens with short working distance to image the vessel. Furthermore, selected agonist and/or antagonists can be utilized to further investigate the mechanisms controlling cell adhesion. Advantages of this method over intravital microscopy include no involvement of invasive surgery and therefore a higher throughput can be obtained. This method also enables the use of localised inhibitor treatment to the desired vessel whereas intravital only enables systemic inhibitor treatment.

This is a method to visualise leukocyte adhesion to the endothelium in harvested pressurised vessels. The technique enables studying vascular adhesion under shear flow with differing intraluminal pressures up to 200 mmHg thus mimic-ing the pathophysiological conditions of high blood pressure.

Worldwide, hypertension is reported to be in approximately a quarter of the population and is the leading biomedical risk factor for mortality worldwide. ...

Immunology and Infection

Long-term Intravital Immunofluorescence Imaging of Tissue Matrix Components with Epifluorescence and Two-photon Microscopy

Besides being a physical scaffold to maintain tissue morphology, the extracellular matrix (ECM) is actively involved in regulating cell and tissue function during development and organ homeostasis. It does so by acting via biochemical, biomechanical, and biophysical signaling pathways, such as through the release of bioactive ECM protein fragments, regulating tissue tension, and providing pathways for cell migration. The extracellular matrix of the tumor microenvironment undergoes substantial remodeling, characterized by the degradation, deposition and organization of fibrillar and non-fibrillar matrix proteins. Stromal stiffening of the tumor microenvironment can promote tumor growth and invasion, and cause remodeling of blood and lymphatic vessels. Live imaging of matrix proteins, however, to this point is limited to fibrillar collagens that can be detected by second harmonic generation using multi-photon microscopy, leaving the majority of matrix components largely invisible. Here we describe procedures for tumor inoculation in the thin dorsal ear skin, immunolabeling of extracellular matrix proteins and intravital imaging of the exposed tissue in live mice using epifluorescence and two-photon microscopy. Our intravital imaging method allows for the direct detection of both fibrillar and non-fibrillar matrix proteins in the context of a growing dermal tumor. We show examples of vessel remodeling caused by local matrix contraction. We also found that fibrillar matrix of the tumor detected with the second harmonic generation is spatially distinct from newly deposited matrix components such as tenascin C. We also showed long-term (12 hours) imaging of T-cell interaction with tumor cells and tumor cells migration along the collagen IV of basement membrane. Taken together, this method uniquely allows for the simultaneous detection of tumor cells, their physical microenvironment and the endogenous tissue immune response over time, which may provide important insights into the mechanisms underlying tumor progression and ultimate success or resistance to therapy.

The extracellular matrix undergoes substantial remodeling during wound healing, inflammation and tumorigenesis. We present a novel intravital immunofluorescence microscopy approach to visualize the dynamics of fibrillar as well as mesh-like matrix components with high spatial and temporal resolution using epifluorescence or two-photon microscopy.

Besides being a physical scaffold to maintain tissue morphology, the extracellular matrix (ECM) is actively involved in regulating cell and tissue ...

Revealing the Cytoskeletal Organization of Invasive Cancer Cells in 3D

Cell migration has traditionally been studied in 2D substrates. However, it has become increasingly evident that there is a need to study cell migration in more appropriate 3D environments, which better resemble the dimensionality of the physiological processes in question. Migratory cells can substantially differ in their morphology and mode of migration depending on whether they are moving on 2D or 3D substrates. Due to technical difficulties and incompatibilities with most standard protocols, structural and functional analysis of cells embedded within 3D matrices still remains uncommon. This article describes methods for preparation and imaging of 3D cancer cell cultures, either as single cells or spheroids. As an appropriate ECM substrate for cancer cell migration, we use nonpepsinized rat tail collagen I polymerized at room-temperature and fluorescently labeled to facilitate visualization using standard confocal microscopes. This work also includes a protocol for 3D immunofluorescent labeling of endogenous cell cytoskeleton. Using these protocols we hope to contribute to a better description of the molecular composition, localization, and functions of cellular structures in 3D.

This article presents a method to fluorescently label collagen that can be further used for both fix and live imaging of 3D cell cultures. We also provide an optimized protocol to visualize endogenous cytoskeletal proteins of cells cultured in 3D environments.

Cell migration has traditionally been studied in 2D substrates. However, it has become increasingly evident that there is a need to study cell migration ...

Medicine

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Historically, most cellular processes have been studied in only 2 dimensions. While these studies have been informative about general cell signaling mechanisms, they neglect important cellular cues received from the structural and mechanical properties of the local microenvironment and extracellular matrix (ECM). To understand how cells interact within a physiological ECM, it is important to study them in the context of 3 dimensional assays. Cell migration, cell differentiation, and cell proliferation are only a few processes that have been shown to be impacted by local changes in the mechanical properties of a 3-dimensional ECM. Collagen I, a core fibrillar component of the ECM, is more than a simple structural element of a tissue. Under normal conditions, mechanical cues from the collagen network direct morphogenesis and maintain cellular structures. In diseased microenvironments, such as the tumor microenvironment, the collagen network is often dramatically remodeled, demonstrating altered composition, enhanced deposition and altered fiber organization. In breast cancer, the degree of fiber alignment is important, as an increase in aligned fibers perpendicular to the tumor boundary has been correlated to poorer patient prognosis1. Aligned collagen matrices result in increased dissemination of tumor cells via persistent migration2,3. The following is a simple protocol for embedding cells within a 3-dimensional, fibrillar collagen hydrogel. This protocol is readily adaptable to many platforms, and can reproducibly generate both aligned and random collagen matrices for investigation of cell migration, cell division, and other cellular processes in a tunable, 3-dimensional, physiological microenvironment.

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Collagen is a core component of the ECM, and provides essential cues for several cellular processes ranging from migration to differentiation and proliferation. Provided here is a protocol for embedding cells within 3D collagen hydrogels, and a more advanced technique for generating randomized or aligned collagen matrices using PDMS microchannels.

Historically, most cellular processes have been studied in only 2 dimensions.  While these studies have been informative about general cell signaling ...

Quantifying Fibrillar Collagen Organization with Curvelet Transform-Based Tools

Fibrillar collagens are prominent extracellular matrix (ECM) components, and their topology changes have been shown to be associated with the progression of a wide range of diseases including breast, ovarian, kidney, and pancreatic cancers. Freely available fiber quantification software tools are mainly focused on the calculation of fiber alignment or orientation, and they are subject to limitations such as the requirement of manual steps, inaccuracy in detection of the fiber edge in noisy background, or lack of localized feature characterization. The collagen fiber quantitation tool described in this protocol is characterized by using an optimal multiscale image representation enabled by curvelet transform (CT). This algorithmic approach allows for the removal of noise from fibrillar collagen images and the enhancement of fiber edges to provide location and orientation information directly from a fiber, rather than using the indirect pixel-wise or window-wise information obtained from other tools. This CT-based framework contains two separate, but linked, packages named &#8220;CT-FIRE&#8221; and &#8220;CurveAlign&#8221; that can quantify fiber organization on a global, region of interest (ROI), or individual fiber basis. This quantification framework has been developed for more than ten years and has now evolved into a comprehensive and user-driven collagen quantification platform. Using this platform, one can measure up to about thirty fiber features including individual fiber properties such as length, angle, width, and straightness, as well as bulk measurements such as density and alignment. Additionally, the user can measure fiber angle relative to manually or automatically segmented boundaries. This platform also provides several additional modules including ones for ROI analysis, automatic boundary creation, and post-processing. Using this platform does not require prior experience of programming or image processing, and it can handle large datasets including hundreds or thousands of images, enabling efficient quantification of collagen fiber organization for biological or biomedical applications.

Here, we present a protocol to use a curvelet transform-based, open-source MATLAB software tool for quantifying fibrillar collagen organization in the extracellular matrix of both normal and diseased tissues. This tool can be applied to images with collagen fibers or other types of line-like structures.

Fibrillar collagens are prominent extracellular matrix (ECM) components, and their topology changes have been shown to be associated with the progression ...

A Method to Study the Correlation Between Local Collagen Structure and Mechanical Properties of Atherosclerotic Plaque Fibrous Tissue

The rupture of atherosclerotic plaques in coronary and carotid arteries is the primary cause of fatal cardiovascular events. However, the rupture mechanics of the heterogeneous, highly collagenous plaque tissue, and how this is related to the tissue's fibrous structure, are not known yet. Existing pipelines to study plaque mechanics are limited to obtaining only gross mechanical characteristics of the plaque tissue, based on the assumption of structural homogeneity of the tissue. However, fibrous plaque tissue is structurally heterogeneous, arguably mainly due to local variation in the collagen fiber architecture.
The mechano-imaging pipeline described here has been developed to study the heterogeneous structural and mechanical plaque properties. In this pipeline, the tissue's local collagen architecture is characterized using multiphoton microscopy (MPM) with second-harmonic generation (SHG), and the tissue's failure behavior is characterized under uniaxial tensile testing conditions using digital image correlation (DIC) analysis. This experimental pipeline enables correlation of the local predominant angle and dispersion of collagen fiber orientation, the rupture behavior, and the strain fingerprints of the fibrous plaque tissue. The obtained knowledge is key to better understand, predict, and prevent atherosclerotic plaque rupture events.

We have developed a mechano-imaging pipeline to study the heterogeneous structural and mechanical atherosclerotic plaque properties. This pipeline enables correlation of the local predominant angle and dispersion of collagen fiber orientation, the rupture behavior, and the strain fingerprints of the fibrous plaque tissue.

The rupture of atherosclerotic plaques in coronary and carotid arteries is the primary cause of fatal cardiovascular events. However, the rupture ...

Dynamic Measurement and Imaging of Capillaries, Arterioles, and Pericytes in Mouse Heart

Coronary arterial tone along with the opening or closing of the capillaries largely determine the blood flow to cardiomyocytes at constant perfusion pressure. However, it is difficult to monitor the dynamic changes of the coronary arterioles and the capillaries in the whole heart, primarily due to its motion and non-stop beating. Here we describe a method that enables monitoring of arterial perfusion rate, pressure and the diameter changes of the arterioles and capillaries in mouse right ventricular papillary muscles. The mouse septal artery is cannulated and perfused at a constant flow or pressure with the other dynamically measured. After perfusion with a fluorescently labeled lectin (e.g., Alexa Fluor-488 or -633 labeled Wheat-Germ Agglutinin, WGA), the arterioles and capillaries (and other vessels) in right ventricle papillary muscle and septum could be readily imaged. The vessel-diameter changes could then be measured in the presence or absence of heart contractions. When genetically encoded fluorescent proteins were expressed, specific features could be monitored. For examples, pericytes were visualized in mouse hearts that expressed NG2-DsRed. This method has provided a useful platform to study the physiological functions of capillary pericytes in heart. It is also suitable for studying the effect of reagents on the blood flow in heart by measuring the vascular/capillary diameter and the arterial luminal pressure simultaneously. This preparation, combined with a state-of-the-art optic imaging system, allows one to study the blood flow and its control at cellular and molecular level in the heart under near-physiological conditions.

Presented here is a protocol to study the coronary microcirculation in living murine heart tissue by ex vivo monitoring of the arterial perfusion pressure and flow that maintains the pressure, as well as vascular tree components including the capillary beds and pericytes, as the septal artery is cannulated and pressurized.

Coronary arterial tone along with the opening or closing of the capillaries largely determine the blood flow to cardiomyocytes at constant perfusion ...

Two-photon Imaging of Intracellular Ca2+ Handling and Nitric Oxide Production in Endothelial and Smooth Muscle Cells of an Isolated Rat Aorta

Calcium is a very important regulator of many physiological processes in vascular tissues. Most endothelial and smooth muscle functions highly depend on changes in intracellular calcium ([Ca2+]i) and nitric oxide (NO). In order to understand how [Ca2+]i, NO and downstream molecules are handled by a blood vessel in response to vasoconstrictors and vasodilators, we developed a novel technique that applies calcium-labeling (or NO-labeling) dyes with two photon microscopy to measure calcium handling (or NO production) in isolated blood vessels. Described here is a detailed step-by-step procedure that demonstrates how to isolate an aorta from a rat, label calcium or NO within the endothelial or smooth muscle cells, and image calcium transients (or NO production) using a two photon microscope following physiological or pharmacological stimuli. The benefits of using the method are multi-fold: 1) it is possible to simultaneously measure calcium transients in both endothelial cells and smooth muscle cells in response to different stimuli; 2) it allows one to image endothelial cells and smooth muscle cells in their native setting; 3) this method is very sensitive to intracellular calcium or NO changes and generates high resolution images for precise measurements; and 4) described approach can be applied to the measurement of other molecules, such as reactive oxygen species. In summary, application of two photon laser emission microscopy to monitor calcium transients and NO production in the endothelial and smooth muscle cells of an isolated blood vessel has provided high quality quantitative data and promoted our understanding of the mechanisms regulating vascular function.

Two-photon Imaging of Intracellular Ca2+ Handling and Nitric Oxide Production in Endothelial and Smooth Muscle Cells of an Isolated Rat Aorta

Vascular cell functiondepends on activity of intracellular messengers. Described here is an ex vivo two photon imaging method that allows the measurement of intracellular calcium and nitric oxide levels in response to physiological and pharmacological stimuli in individual endothelial and smooth muscle cells of an isolated aorta.

Calcium is a very important regulator of many physiological processes in vascular tissues. Most endothelial and smooth muscle functions highly depend on ...

Assessing Collagen and Elastin Pressure-dependent Microarchitectures in Live, Human Resistance Arteries by Label-free Fluorescence Microscopy

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