We thank Katie Carroll for technical assistance. We also thank Dr. David Cool and the Proteomics Analysis Laboratory for instruction on and provision of the Cytation imaging plate reader. This work was supported by a Wright State Foundation grant (to L.E.W.).

1. Detection of EdU using fluorescent label
<ol>
	<li>Stock solutions
	<ol>
		<li>Prepare a 5 mM EdU stock solution by adding 12.5 mg of EdU to 10 mL of double distilled H2O.</li>
		<li>Prepare the labeling solution components: 200 mM tris(3-hydroxypropyltriazolylmethyl)amine (THPTA) (100 mg in 1.15 mL H2O), 100 mM CuSO4 (15.95 mg in 1 mL H2O; make fresh), 10 mM Cy3 picolyl-azide (1 mg in 95.3 mL H2O, stored in 5 &#181;L aliquots at -20 &#176;C), and 1 M sodium ascorbate (200 mg in 1 mL H2O; make fresh).</li>
		<li>Prepare resazurin stock solution at a concentration of 0.15 mg/mL. Filter sterilize and store 1 mL aliquots at 4 &#176;C. 
		NOTE: Picolyl azide is available conjugated to several different fluors, biotin, and horseradish peroxidase (HRP).</li>
	</ol>
	</li>
	<li>Label vascular smooth muscle cells (VSMC) with EdU. 
	NOTE: Human VSMCs were isolated from iliac arteries via outgrowth from explanted pieces of tissue. When grown in serum free media with insulin, transferrin, selenium these cells express VSMC markers smoothelin, smooth muscle myosin heavy chain (SM-MHC) and SMC &#945;actin13.
	<ol>
		<li>Plate vascular smooth muscle cells at 2 x 104/mL in Dulbecco&#8217;s modified Eagle&#8217;s medium (DMEM) in a 96 well plate. 
		NOTE: Cells are grown at 37 &#176;C in smooth muscle cell media with 2% fetal bovine serum.</li>
		<li>(Optional) Allow several hours to overnight for adherence. Add 20 &#181;L resazurin stock to each well. Incubate at 37 &#176;C for 3 h. Read the fluorescence signal in a plate reader (560ex, 594em). Replace media with fresh DMEM. 
		NOTE: This step is optional. Resazurin is used to normalize cell numbers and account for well-to-well variability in plating14.</li>
		<li>Add phosphate buffered saline (PBS) or platelet derived growth factor (PDGF) 30 ng/mL to 3&#8722;6 wells per treatment at the beginning of the culture period and incubate for 72 h.</li>
		<li>Dilute 5 mM EdU stock to 1 mM. Add 2 &#181;L to each well (final concentration 20 &#181;M) for the last 24 h of the 72 h culture period. Do not add EdU to one set of replicates. These wells will be used to determine background fluorescence/luminescence. 
		NOTE: The duration of culture and labeling with EdU is optimized for smooth muscle cells but may need to be modified per cell type.</li>
	</ol>
	</li>
	<li>Fix cells in paraformaldehyde (PFA).
	<ol>
		<li>Remove media and add 150 &#181;L of 4% PFA (in PBS) per well for 10 min at room temperature.</li>
		<li>Remove PFA and add 150 &#181;L of 1% polyethylene glycol&#160;tert-octylphenyl ether (TX-100) (in water) per well for 30 min at room temperature.</li>
		<li>Remove TX-100 by washing three times with PBS. 
		CAUTION: PFA is toxic, handle with care. 
		NOTE: The protocol can be paused here. Fixed cells can be stored in PBS at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Detect incorporated EdU.
	<ol>
		<li>Make labeling solution (10 mL per 96 well plate, using inner 60 wells) just prior to use by adding reagents from stock solutions in the following order: THPTA 20 &#181;L, CuSO4 20 &#181;L, Cy3 picolyl azide 5 &#181;L, Na ascorbate 100 &#181;L to PBS for a total volume of 10 mL.</li>
		<li>Remove PBS from wells and add 150 &#181;L of labeling solution to each well. Incubate 30 min at 37 &#176;C. To detect all nuclei, add 4&#8242;,6-diamidino-2-phenylindole (DAPI) to each well. Read on a fluorescence plate reader (Cy3 550ex, 570em; DAPI 350ex, 470em) or image on a microscope. 
		NOTE: The protocol can be paused here. Fixed and stained cells can be stored in PBS at 4 &#176;C and imaged at a later time.</li>
	</ol>
	</li>
</ol>
2. Detection of EdU using luminescence
<ol>
	<li>Complete sections 1.1&#8211;1.3.</li>
	<li>Prepare Tris-buffered saline with 0.1% polysorbate 20 (TBST).</li>
	<li>Block wells.
	<ol>
		<li>Add 150 &#181;L of blocking buffer (Table of Materials) to each well and incubate for 90 min at room temperature.</li>
		<li>Remove blocking buffer and wash three times with 200 &#181;L of PBS.</li>
	</ol>
	</li>
	<li>Detect incorporated EdU.
	<ol>
		<li>Make labeling solution as directed in step 1.4.1, substituting biotin picolyl azide for Cy3 picolyl azide.</li>
		<li>Wash wells five times with 200 &#181;L of TBST, with shaking, 3 min per wash.</li>
		<li>Quench endogenous peroxidases with 150 &#181;L of 0.3% H2O2 for 20 min at room temperature.</li>
		<li>Remove H2O2 and wash five times with 200 &#181;L of TBST, with shaking 3 min per wash.</li>
		<li>Dilute streptavidin-horseradish peroxidase (SA-HRP; 1:200) in TBST and add 50 &#181;L to each well. Incubate 1 h at room temperature.</li>
		<li>Wash five times with 200 &#181;L of TBST, with shaking, 3 min per wash.</li>
		<li>Add 50 &#181;L of chemiluminescent enzyme-linked immunosorbent assay (ELISA) substrate (Table of Materials) to each well.</li>
		<li>Read immediately in plate reader capable of detecting luminescence.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Incorporation of EdU is a simple, straightforward way to measure cell proliferation; it is particularly useful for adherent cells13. Our protocol uses smooth muscle cells, but it is applicable to any adherent cell (epithelial, endothelial, etc.). Although the protocol is not complicated, the one critical step is making the labeling solution &#8212; the ingredients must be added in the order listed. In addition, like chemiluminescent Western blots, if using the luminescence readout the plate must be read immediately.
Several parts of the protocol can be modified or optimized as needed. One part is the duration of incubation with EdU. Because our goal was to detect as many proliferating cells as possible, we added EdU for a full 24 h prior to fixing and staining. However, this timing likely measures cells in both S and G2/M phases. To label only cells in S phase, Cecchini et al. advise that even slowly dividing cells be incubated with EdU for no longer than 6 hours7. Another part of the protocol that can be optimized is the amount of chelator. Alkyne-azide reactions require a copper catalyst (CuSO4). THPTA is a copper chelator that maintains copper in the necessary oxidation state. The ratio of chelator to copper can be varied, to increase alkyne-azide binding. We observe bright staining with a chelator:copper ratio of 2:1, in the context of also using picolyl azide. The picolyl moiety adds chelating activity which decreases the concentration of copper needed and increases the efficiency of the reaction15. Chelator:copper ratios can be increased, usually up to 5:1, to optimize per investigator needs. A better chelator, 2-[4-({bis[(1-tert-butyl-1H-1,2,3-triazol-4-yl)methyl]amino}methyl)-1H-1,2,3-triazol-1-yl]acetic acid (BTTAA), is now commercially available16. This chelator was not commercially available when we first derived this protocol. Finally, although we use paraformaldehyde as a fixative in this protocol, cells can be fixed with methanol, which also obviates the permeabilization step. Fixing with methanol can be helpful if one wants to combine immunofluorescent staining of specific cellular proteins with incorporation of EdU.
One drawback of this method is the potential cytotoxicity of EdU, particularly when continuously labeling with EdU for days as opposed to a pulse label of hours. Depending on the cells used, EdU has been found to cause cell cycle arrest or even apoptosis17. This toxicity is thought to be due to defective DNA replication from the addition of artificial nucleosides18. Studies have shown this toxicity to be cell line dependent and concentration dependent17. We did not observe any problems with toxicity in our studies.
In summary, measuring proliferation via EdU and click chemistry has several advantages over other commonly used methods such as BrdU or 3H-thymidine incorporation. These include avoiding radioactivity, high specificity with low background, and no need for harsh denaturation steps. Once established, click chemistry is highly versatile, and can be easily modified for the metabolic labeling of RNA, protein, fatty acids, and carbohydrates.

Proliferation is a critical part of cellular function1,2. Control of proliferation influences normal processes such as development, and pathologic processes such as cancer and cardiovascular disease. Hyperplastic growth of vascular smooth muscle cells, for example, is thought to be a precursor to atherosclerosis3. Changes in cell proliferation are also used to assess potential toxicity of new drugs. Given its widespread impact, measuring proliferation is a mainstay of many cell biology-based laboratories.
Cell proliferation can be measured by simply counting cells if the cell population of interest divides fairly rapidly. For slower growing cells, cell counts may be less sensitive. Proliferation is often measured by incorporation of labeled nucleoside analogs. Although the gold standard is 3H-thymidine incorporation, many laboratories are getting away from this method given the availability of newer, non-radioactive alternatives. These include cytoplasmic fluorescent dyes, detection of cell cycle associated proteins, and incorporation of non-radioactive nucleoside analogs such as 5-bromo-2&#8217; deoxyuridine (BrdU) and 5-ethynyl-2&#8242;-deoxyuridine (EdU)4.
Cytoplasmic dyes, such as carboxyfluorescein diacetate succinimidyl ester (CFSE), detect proliferation because the intensity of the dye halves each time the cell divides5. This technique is commonly used in flow cytometry for non-adherent cells. It has not been used much for adherent cells, but with the new generation of imaging plate readers this may change. Detection of cell cycle associated proteins through antibody-based techniques (flow cytometry, immunohistochemistry, etc.) is often used for tissues or non-adherent cells. These cells/tissues must be fixed and permeabilized prior to staining. Nucleoside analogs BrdU and EdU are similar in approach to 3H-thymidine, but without the inconvenience of radioactivity. Incorporation of BrdU is detected by anti-BrdU antibodies, and therefore cells must be fixed and permeabilized prior to staining. In addition, cells must be treated with DNase to expose the BrdU epitope.
Incorporation of EdU is detected by click chemistry, in which alkyne and azide groups &#8220;click&#8221; together in the presence of catalytic copper. Either group can serve as the biosynthetically incorporated molecule or detection molecule4. Commercially available nucleoside analogs have the alkyne group attached. For proliferation assays, therefore, the alkyne functionalized nucleoside analog is detected by an azide conjugated to a fluorescent dye or other marker. Incorporation of EdU has several advantages over BrdU. First, because alkyne and azide groups are not found in mammalian cells, this interaction is highly specific with a low background6. Because both groups are small, cells do not need to undergo DNA denaturation to expose the nucleoside analog as required for BrdU7. Finally, click chemistry is highly versatile, and can be used for the metabolic labeling of DNA, RNA, protein, fatty acids, and carbohydrates8,9,10,11. If the metabolically labeled target of interest is on the cell surface, live cells can be labeled12. In addition, cells can be further processed for immunofluorescent staining with antibodies.
The following protocol describes the use of EdU incorporation and click chemistry to measure proliferation in human vascular smooth muscle cells (Figure 1). We show three different ways incorporation of EdU can be measured, representative results from each, and their advantages and disadvantages. Measuring proliferation via click chemistry is particularly useful for adherent, slower growing cells. In addition, the morphology of the cell is well maintained and antibody-based staining can also be performed.

<table><tbody><tr><td>5-Ethynyl-2&#039;-deoxyuridine</td><td>Carbosynth</td><td>NE08701</td><td></td></tr><tr><td>biotin picolyl azide</td><td>Click Chemistry Tools</td><td>1167-5</td><td></td></tr><tr><td>CuSO4</td><td>Fisher Scientific</td><td>C1297-100G</td><td></td></tr><tr><td>Cy3 picolyl azide</td><td>Click Chemistry Tools</td><td>1178-1</td><td></td></tr><tr><td>Cytation Plate Reader</td><td>Biotek</td><td></td><td></td></tr><tr><td>EVOS Microscope</td><td>Thermo Fisher Scientific</td><td></td><td></td></tr><tr><td>Hydrogen peroxide solution</td><td>Sigma-Aldrich</td><td>216763</td><td></td></tr><tr><td>Na ascorbate</td><td>Sigma-Aldrich</td><td>11140-250G</td><td></td></tr><tr><td>NucBlue Fixed Cell Stain Ready Probes</td><td>Invitrogen</td><td>R37606</td><td>DAPI nuclear stain</td></tr><tr><td>Odyssey Blocking Buffer</td><td>Li-Cor</td><td>927-40000</td><td>blocking buffer</td></tr><tr><td>Paraformaldehyde</td><td>Electron Microscopy Services</td><td>15710</td><td></td></tr><tr><td>PBS</td><td>Fisher Scientific</td><td>SH3002802</td><td></td></tr><tr><td>Sodium Chloride</td><td>Sigma Life Science</td><td>S3014-5kg</td><td></td></tr><tr><td>Streptavidin Horseradish Peroxidase Conjugate</td><td>Life Technologies</td><td>S911</td><td></td></tr><tr><td>Super Signal ELISA Femto</td><td>Thermo Scientific</td><td>PI137075</td><td>ELISA substrate</td></tr><tr><td>Synergy Plate Reader</td><td>Biotek</td><td></td><td></td></tr><tr><td>THPTA</td><td>Click Chemistry Tools</td><td>1010-100</td><td></td></tr><tr><td>Tris Hydroxymethyl Aminomethane Hydrochloride (Tris-HCl)</td><td>Fisher Scientific</td><td>BP153-500</td><td></td></tr><tr><td>Triton X 100</td><td>Bio-Rad</td><td>1610407</td><td></td></tr><tr><td>Tween 20</td><td>Fisher Scientific</td><td>BP337-100</td><td></td></tr></tbody></table>

measuring proliferation of vascular smooth muscle cells using click chemistry

The ability of a cell to proliferate is integral to the normal function of most cells, and dysregulation of proliferation is at the heart of many disease processes. For these reasons, measuring proliferation is a common tool used to assess cell function. Cell proliferation can be measured simply by counting; however, this is an indirect means of measuring proliferation. One common means of directly detecting cells preparing to divide is by incorporation of labeled nucleoside analogs. These include the radioactive nucleoside analog 3H-thymidine plus non-radioactive nucleoside analogs such as 5-bromo-2&#8217; deoxyuridine (BrdU) and 5-ethynyl-2&#8242;-deoxyuridine (EdU). Incorporation of EdU is detected by click chemistry, which has several advantages when compared to BrdU. In this report, we provide a protocol for measuring proliferation by the incorporation of EdU. This protocol includes options for various readouts, along with the advantages and disadvantages of each. We also discuss places where the protocol can be optimized or altered to meet the specific needs of the experiment planned. Finally, we touch on the ways that this basic protocol can be modified for measuring other cell metabolites.

In Figure 2, we demonstrate the outcomes of three different experiments measuring proliferation of VSMC in response to PDGF. After growing the cells in media formulated for SMCs, we carried out the experiment in serum free DMEM, to eliminate any potential effects of serum or growth factors on proliferation. In Figure 2A we compare results, from the same experiment, using a fluorescent vs luminescent readout. To read these plates, we used a multimode microplate reader that can read absorbance, fluorescence, and luminescence (see Table of Materials).
Using the plate reader in scanning well mode, fluorescent readings are averaged over the entire well. This mode alleviates potential inaccuracies due to differences between the center and edges of the well. However, if very few nuclei are positive, reading results in this way likely underestimates the &#8220;number&#8221; of positive cells when only a few are positive. Number, with this readout, means fluorescence relative to that of another well, and not exact cell numbers. In addition, if there is a fiber or a clump of dead cells that picks up fluorescence, this will be included in the area scan. However, the advantage of using the plate reader is time. While reading in scanning mode takes 15&#8722;20 seconds per well, it is automated as opposed to the personal time-intensive method of taking pictures and analyzing them using imaging software.
To correct potential underestimates from the above method, we converted the fluorescent scan to a homogenous, liquid readout in which cells taking up EdU are detected with an azide-biotin moiety, which is in turn detected with streptavidin-horseradish peroxidase (see Figure 1). The amount of horseradish peroxidase is then quantified using a standard substrate formatted for ELISA. The advantage of this technique is a more homogenous readout, and Figure 2A,B shows examples of experiments where this method was employed. In addition, the plate can be read in seconds. There are disadvantages to this method, however. First, the background with this method is higher than with fluorescence. To decrease the background, we incorporated a blocking step and multiple washes. As a reflection of this background, negative controls (untreated cells) tend to be higher, and the fold increase in proliferation is lower. Second, as with the fluorescence readout, any fibers or dead cells that pick up the detection reagents will add to the total luminescence.
In an attempt to decrease variability in the above experiments, we normalized our results to initial cell counts. For proliferation assays, normalization has to be done on live cells at the beginning of the assay before the cells divide. In addition, the assay cannot affect cell function or incorporation of EdU. Given these constraints, we chose to use metabolism as a reflection of cell number via resazurin14. Results with and without normalization are shown in Figure 2B. However, in a separate set of experiments we found that this method is not sensitive enough to detect small differences in cell numbers. In addition, any error in this readout introduces more error in the entire experiment. For these reasons, we have elected to take extraordinary care to plate cells as evenly as possible, and not perform a normalization step.
To be sure we were getting the most accurate results possible, we reverted to fluorescent staining (section 1) and counting cells using an inverted fluorescence microscope. The advantage of this method is that one can physically see and count the positive cells, ensuring the most accuracy (Figure 3). Any fluorescent debris can be excluded from the count. Background counts without EdU are 0 as there is no visible fluorescence, and therefore no background to subtract. The main disadvantage of this method is time. To count nuclei in cells plated in a 96 well plate, we take 3 pictures per well of both EdU positive cells and total cells (DAPI), requiring 6 images per well. We take pictures vertically from the midsection of the top to the bottom of the well, taking care not to overlap and count cells twice. Counting total cells gives us a second piece of data that confirms our proliferation results. However, in light of the relatively slow doubling time of VSMC (36&#8722;48 h), the fold increase in total cells stimulated with PDGF vs controls is much lower than the fold increase of EdU positive cells, which is why we measure incorporation of EdU (Figure 2C).
The most efficient and accurate way to count EdU positive and total cells is by using an imaging plate reader. This method combines the accuracy of counting with the speed of automation. The main drawback is that this piece of equipment is not available at every institution. In comparing automated vs manual counts, the EdU positive manual count was essentially identical to the automated count (Figure 2C, left), as were the total unstimulated counts (Figure 2C, right). However, the manual count was lower than the automated count in total PDGF-stimulated cells. This difference likely relates to cell growth patterns. Because the cells tend to grow in the center of the well, with smaller numbers the automated and manual counts were nearly identical. However, with higher numbers there were likely more cells at the edges and the manual counts, which did not capture cells at the periphery of the well, were less accurate.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59930/59930fig01.jpg" /> 
Figure 1: Overview of EdU detection by fluorescence or luminescence. 
VSMC or other adherent cells are treated under conditions of interest. EdU is added for the last 24 h of the culture period and incorporated into the DNA of dividing cells. Incorporated EdU is then detected using fluorescence (protocol section 1) or luminescence (protocol section 2). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/59930/59930fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59930/59930fig02.jpg" /> 
Figure 2: Measuring proliferation with different readouts using incorporation of EdU and click chemistry. 
(A) VSMCs were cultured in the presence or absence of PDGF. EdU was added the last 24 h of culture, and incorporation was detected by either fluorescence (protocol section 1) or luminescence (protocol section 2). (B) VSMCs were treated as described in panel A, and incorporation of EdU was measured by luminescence. Results were normalized to initial cell counts using resazurin viability assay. (C) VSMCs were treated as in panel A, and incorporation of EdU was measured by fluorescence. EdU positive cells were counted automatically, using an imaging plate reader, or manually by imaging and then counting using imaging software. Results shown are the average &#177; standard deviation of 3&#8211;6 replicates per treatment.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59930/59930fig03.jpg" /> 
Figure 3: Visualization of EdU positive cells following click fluorescence protocol. 
VSMC were treated as described in Figure 2, and incorporated EdU was detected using fluorescence (protocol section 1). Each image is a single well of a 96 well plate. Total nuclei are shown in blue (DAPI) and EdU positive nuclei are green. Scale bar = 50 &#181;m. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/59930/59930fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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Measuring Proliferation of Vascular Smooth Muscle Cells Using Click Chemistry

Proliferation is a critical part of cellular function, and a common readout used to assess potential toxicity of new drugs. Measuring proliferation is, therefore, a frequently used assay in cell biology. Here we present a simple, versatile method of measuring proliferation that can be used in adherent and non-adherent cells.

The ability of a cell to proliferate is integral to the normal function of most cells, and dysregulation of proliferation is at the heart of many disease ...

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Universidad San Sebastian

Research

JoVE Journal

Biology

6.7K Views.  Wright State University. Proliferation is a critical part of cellular function, and a common readout used to assess potential toxicity of new drugs. Measuring proliferation is, therefore, a frequently used assay in cell biology. Here we present a simple, versatile method of measuring proliferation that can be used in adherent and non-adherent cells.

Video: Measuring Proliferation of Vascular Smooth Muscle Cells Using Click Chemistry

Assessment of Endothelial Cell Migration After Exposure to Toxic Chemicals

Exposure to chemical substances (including alkylating chemical warfare agents like sulfur and nitrogen mustards) cause a plethora of clinical symptoms including wound healing disorder. The physiological process of wound healing is highly complex. The formation of granulation tissue is a key step in this process resulting in a preliminary wound closure and providing a network of new capillary blood vessels &#8211; either through vasculogenesis (novel formation) or angiogenesis (sprouting of existing vessels). Both vasculo- and angiogenesis require functional, directed migration of endothelial cells. Thus, investigation of early endothelial cell (EEC) migration is important to understand the pathophysiology of chemical induced wound healing disorders and to potentially identify novel strategies for therapeutic intervention.
We assessed impaired wound healing after alkylating agent exposure and tested potential candidate compounds for treatment. We used a set of techniques outlined in this protocol. A modified Boyden chamber to quantitatively investigate chemokinesis of EEC is described. Moreover, the use of the wound healing assay in combination with track analysis to qualitatively assess migration is illustrated. Finally, we demonstrate the use of the fluorescent dye TMRM for the investigation of mitochondrial membrane potential to identify underlying mechanisms of disturbed cell migration. The following protocol describes basic techniques that have been adapted for the investigation of EEC.

Investigation of early endothelial cell (EEC) migration is important to understand the pathophysiology of certain illnesses and to potentially identify novel strategies for therapeutic intervention. The following protocol describes techniques to assess cell migration that have been adapted for the investigation of EEC.

Exposure to chemical substances (including alkylating chemical warfare agents like sulfur and nitrogen mustards) cause a plethora of clinical symptoms ...

Developmental Biology

Electric Cell-substrate Impedance Sensing for the Quantification of Endothelial Proliferation, Barrier Function, and Motility

Electric Cell-substrate Impedance Sensing (ECIS) is an in vitro impedance measuring system to quantify the behavior of cells within adherent cell layers. To this end, cells are grown in special culture chambers on top of opposing, circular gold electrodes. A constant small alternating current is applied between the electrodes and the potential across is measured. The insulating properties of the cell membrane create a resistance towards the electrical current flow resulting in an increased electrical potential between the electrodes. Measuring cellular impedance in this manner allows the automated study of cell attachment, growth, morphology, function, and motility. Although the ECIS measurement itself is straightforward and easy to learn, the underlying theory is complex and selection of the right settings and correct analysis and interpretation of the data is not self-evident. Yet, a clear protocol describing the individual steps from the experimental design to preparation, realization, and analysis of the experiment is not available. In this article the basic measurement principle as well as possible applications, experimental considerations, advantages and limitations of the ECIS system are discussed. A guide is provided for the study of cell attachment, spreading and proliferation; quantification of cell behavior in a confluent layer, with regard to barrier function, cell motility, quality of cell-cell and cell-substrate adhesions; and quantification of wound healing and cellular responses to vasoactive stimuli. Representative results are discussed based on human microvascular (MVEC) and human umbilical vein endothelial cells (HUVEC), but are applicable to all adherent growing cells.

This protocol reviews Electric Cell-substrate Impedance Sensing, a method to record and analyze the impedance spectrum of adherent cells for the quantification of cell attachment, proliferation, motility, and cellular responses to pharmacological and toxic stimuli. Detection of endothelial permeability and assessment of cell-cell and cell-substrate contacts are emphasized.

Electric Cell-substrate Impedance Sensing (ECIS) is an in vitro impedance measuring system to quantify the behavior of cells within adherent cell layers. ...

Bioengineering

Optimized Incorporation of Alkynyl Fatty Acid Analogs for the Detection of Fatty Acylated Proteins using Click Chemistry

Fatty acylation, the covalent addition of saturated fatty acids to protein substrates, is important in regulating a myriad of cellular functions in addition to its implications in cancer and neurodegenerative diseases. Recent developments in fatty acylation detection methods have enabled efficient and non-hazardous detection of fatty acylated proteins, particularly through the use of click chemistry with bio-orthogonal labeling. However, click chemistry detection can be limited by the poor solubility and potential toxic effects of adding long chain fatty acids to cell culture. Described here is a labeling approach with optimized delivery using saponified fatty acids in combination with fatty-acid free BSA, as well as delipidated media, which can improve detection of hard to detect fatty acylated proteins. This effect was most pronounced with the alkynyl-stearate analog, 17-ODYA, which has been the most commonly used fatty acid analog in click chemistry detection of acylated proteins. This modification will improve cellular incorporation and increase sensitivity to acylated protein detection. In addition, this approach can be applied in a variety of cell types and combined with other assays such as pulse-chase analysis, stable isotope labeling with amino acids in cell culture, and mass spectrometry for quantitative profiling of fatty acylated proteins.

The study of fatty acylation has important implications in cellular protein interactions and diseases. Presented here is a modified protocol to improve click chemistry detection of fatty acylated proteins, which can be applied in various cell types and combined with other assays, including pulse-chase and mass spectrometry.

Fatty acylation, the covalent addition of saturated fatty acids to protein substrates, is important in regulating a myriad of cellular functions in ...

Biochemistry

Micropatterning and Assembly of 3D Microvessels

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass based substrates, and hydrogel-based tube formation assays. These assays, while informative, do not recapitulate lumen geometry, proper extracellular matrix, and multi-cellular proximity, which play key roles in modulating vascular function. This manuscript describes an injection molding method to generate engineered vessels with diameters on the order of 100 &#181;m. Microvessels are fabricated by seeding endothelial cells in a microfluidic channel embedded within a native type I collagen hydrogel. By incorporating parenchymal cells within the collagen matrix prior to channel formation, specific tissue microenvironments can be modeled and studied. Additional modulations of hydrodynamic properties and media composition allow for control of complex vascular function within the desired microenvironment. This platform allows for the study of perivascular cell recruitment, blood-endothelium interactions, flow response, and tissue-microvascular interactions. Engineered microvessels offer the ability to isolate the influence from individual components of a vascular niche and precisely control its chemical, mechanical, and biological properties to study vascular biology in both health and disease.

This manuscript presents an injection molding method to engineer microvessels that recapitulate physiological properties of endothelium. The microfluidic-based process creates patent 3D vascular networks with tailorable conditions, such as flow, cellular composition, geometry, and biochemical gradients. The fabrication process and examples of potential applications are described.

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass ...

Monitoring Functionality and Morphology of Vasculature Recruited by Factors Secreted by Fast-growing Tumor-generating Cells

The subcutaneous matrigel plug assay in mice is a method of choice for the in vivo evaluation of pro- and anti-angiogenic factors. In this method, desired factors are introduced into cold-liquid ECM-mimic gel which, after subcutaneous injection, solidifies to form an environment mimicking the cancer milieu. This matrix permits the penetration of host cells, such as endothelial cells, and therefore, the formation of vasculature.
Herein we propose a new modified matrigel plug assay, which can be exploited to illustrate the angiogenic potential of a pool of factors secreted by cancer cells, as opposed to a specific factor (e.g., bFGF and VEGF) or agent. The plug containing ECM-mimic gel is utilized to introduce the host (i.e., mouse) with a pool of factors secreted to the C.M. of fast-growing tumor-generating glioblastoma cells. We have previously described an extensive comparison of the angiogenic potential of U-87 MG human glioblastoma and its dormant-derived clone, in this system model, showing induced angiogenesis in the U-87 MG parental cells. The C.M. is prepared by filtering collected media from confluent tissue culture plates of either cell line following 48 hr incubation. Hence, it contains only factors secreted by the cells, without the cells themselves. Described here is the combination of two imaging modalities, microbubbles contrast-enhanced ultrasound imaging and intravital fibered-confocal endomicroscopy, for an accurate, real-time characterization of the extent, morphology and functionality of newly-formed blood vessels within the plugs.

We describe a matrigel plug assay to illustrate angiogenic potential of a pool of factors secreted by cancer cells using two complementary imaging modalities, ultrasound and endomicroscopy. The matrigel, an extracellular matrix (ECM)-mimic gel, is utilized to introduce the host (mouse) with angiogenic factors secreted to the conditioned media (C.M.).&#160;

The subcutaneous matrigel plug assay in mice is a method of choice for the in vivo evaluation of pro- and anti-angiogenic factors. In this method, desired ...

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 ...

Quantitation of Endothelial Cell Adhesiveness In Vitro

One of the cardinal processes of inflammation is the infiltration of immune cells from the lumen of the blood vessel to the surrounding tissue. This occurs when endothelial cells, which line blood vessels, become adhesive to circulating immune cells such as monocytes. In vitro measurement of this adhesiveness has until now been done by quantifying the total number of monocytes that adhere to an endothelial layer either as a direct count or by indirect measurement of the fluorescence of adherent monocytes. While such measurements do indicate the average adhesiveness of the endothelial cell population, they are confounded by a number of factors, such as cell number, and do not reveal the proportion of endothelial cells that are actually adhesive. Here we describe and demonstrate a method which allows the enumeration of adhesive cells within a tested population of endothelial monolayer. Endothelial cells are grown on glass coverslips and following desired treatment are challenged with monocytes (that may be fluorescently labeled). After incubation, a rinsing procedure, involving multiple rounds of immersion and draining, the cells are fixed. Adhesive endothelial cells, which are surrounded by monocytes are readily identified and enumerated, giving an adhesion index that reveals the actual proportion of endothelial cells within the population that are adhesive.

Quantitation of Endothelial Cell Adhesiveness In Vitro

We report an in vitro method that allows the quantitation of the actual number of adhesive cells within an endothelial cell monolayer.

One of the cardinal processes of inflammation is the infiltration of immune cells from the lumen of the blood vessel to the surrounding tissue. This ...

In Vivo Study of Human Endothelial-Pericyte Interaction Using the Matrix Gel Plug Assay in Mouse

Angiogenesis is the process by which new blood vessels are formed from existing vessels. New vessel growth requires coordinated endothelial cell proliferation, migration, and alignment to form tubular structures followed by recruitment of pericytes to provide mural support and facilitate vessel maturation. Current in vitro cell culture approaches cannot fully reproduce the complex biological environment where endothelial cells and pericytes interact to produce functional vessels. We present a novel application of the in vivo matrix gel plug assay to study endothelial-pericyte interactions and formation of functional blood vessels using severe combined immune deficiency mutation (SCID) mice. Briefly, matrix gel is mixed with a solution containing endothelial cells with or without pericytes followed by injection into the back of anesthetized SCID mice. After 14 days, the matrix gel plugs are removed, fixed and sectioned for histological analysis. The length, number, size and extent of pericyte coverage of mature vessels (defined by the presence of red blood cells in the lumen) can be quantified and compared between experimental groups using commercial statistical platforms. Beyond its use as an angiogenesis assay, this matrix gel plug assay can be used to conduct genetic studies and as a platform for drug discovery. In conclusion, this protocol will allow researchers to complement available in vitro assays for the study of endothelial-pericyte interactions and their relevance to either systemic or pulmonary angiogenesis.

In Vivo Study of Human Endothelial-Pericyte Interaction Using the Matrix Gel Plug Assay in Mouse

We present a protocol to study human endothelial-pericyte interactions in mouse using a variation of the matrix gel plug angiogenesis assay.

Angiogenesis is the process by which new blood vessels are formed from existing vessels. New vessel growth requires coordinated endothelial cell ...

Utilizing the Precision-Cut Lung Slice to Study the Contractile Regulation of Airway and Intrapulmonary Arterial Smooth Muscle

Smooth muscle cells (SMC) mediate the contraction of the airway and the intrapulmonary artery to modify airflow resistance and pulmonary circulation, respectively, hence playing a critical role in the homeostasis of the pulmonary system. Deregulation of SMC contractility contributes to several pulmonary diseases, including asthma and pulmonary hypertension. However, due to limited tissue access and a lack of culture systems to maintain in vivo SMC phenotypes, molecular mechanisms underlying the deregulated SMC contractility in these diseases remain fully identified. The precision-cut lung slice (PCLS) offers an ex vivo model that circumvents these technical difficulties. As a live, thin lung tissue section, the PCLS retains SMC in natural surroundings and allows in situ tracking of SMC contraction and intracellular Ca2+ signaling that regulates SMC contractility. Here, a detailed mouse PCLS preparation protocol is provided, which preserves intact airways and intrapulmonary arteries. This protocol involves two essential steps before subjecting the lung lobe to slicing: inflating the airway with low-melting-point agarose through the trachea and infilling pulmonary vessels with gelatin through the right ventricle. The PCLS prepared using this protocol can be used for bioassays to evaluate Ca2+-mediated contractile regulation of SMC in both the airway and the intrapulmonary arterial compartments. When applied to mouse models of respiratory diseases, this protocol enables the functional investigation of SMC, thereby providing insight into the underlying mechanism of SMC contractility deregulation in diseases.

The present protocol describes preparing and utilizing mouse precision-cut lung slices to assess the airway and intrapulmonary arterial smooth muscle contractility in a nearly in vivo milieu.

Smooth muscle cells (SMC) mediate the contraction of the airway and the intrapulmonary artery to modify airflow resistance and pulmonary circulation, ...

Bromodeoxyuridine Pulse Labelling Assay: A Technique to Measure Cell Proliferation

Source: Welschinger R. et al. <a href="https://www-jove-com.bdigitaluss.remotexs.co/t/52840">Temporal Tracking of Cell Cycle Progression Using Flow Cytometry without the Need for Synchronization.</a> J. Vis. Exp. (2015)
The video describes the stepwise pulse labeling assay of mammalian cells with bromodeoxyuridine (BrdU) for measuring cell proliferation. The BrdU uptake permits the temporal tracking of cells that were in the synthesis phase at a specific point in time, without requiring the cells to be synchronized.

Source: Welschinger R. et al. Temporal Tracking of Cell Cycle Progression Using Flow Cytometry without the Need for Synchronization. J. Vis. Exp. (2015)
...

Measuring Proliferation of Vascular Smooth Muscle Cells Using Click Chemistry

Summary

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Chapters in this video

Assessment of Endothelial Cell Migration After Exposure to Toxic Chemicals

Electric Cell-substrate Impedance Sensing for the Quantification of Endothelial Proliferation, Barrier Function, and Motility

Optimized Incorporation of Alkynyl Fatty Acid Analogs for the Detection of Fatty Acylated Proteins using Click Chemistry

Micropatterning and Assembly of 3D Microvessels

Monitoring Functionality and Morphology of Vasculature Recruited by Factors Secreted by Fast-growing Tumor-generating Cells

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

Quantitation of Endothelial Cell Adhesiveness In Vitro

In Vivo Study of Human Endothelial-Pericyte Interaction Using the Matrix Gel Plug Assay in Mouse

Utilizing the Precision-Cut Lung Slice to Study the Contractile Regulation of Airway and Intrapulmonary Arterial Smooth Muscle

Bromodeoxyuridine Pulse Labelling Assay: A Technique to Measure Cell Proliferation