The authors thank Libby Perry (Augusta University) for her help with the SEM sample preparation. This work was supported by the National Institutes of Health [R01HL139562 (G.C.) and K99HL146954 (B.S.)] and American Heart Association [17POST33661254 (B.S.)].

NOTE: The following is a general protocol used to quantify membrane ruffle formation in RAW 264.7 macrophages using SEM microscopy. Optimization may be required for different cell types.
1. Cell line and cell culture
<ol>
	<li>Grow RAW 264.7 macrophages in Dulbecco&#39;s Modified Eagle Medium (DMEM) supplemented with 10% (vol/vol) heat-inactivated Fetal Bovine Serum (FBS), 100 IU/mL of penicillin and 100 &#181;g/mL streptomycin in a humidified incubator at 37 &#176;C and 5% CO2. Replace growth media every other day.</li>
	<li>When cells become 80% confluent, wash the plate two times using sterile PBS.</li>
	<li>Detach cells by adding 1,000 &#181;L of 0.5% trypsin-EDTA and incubate the cell culture plate for 3-5 min in a humidified incubator at 37 &#176;C.</li>
	<li>Examine the plate under a light microscope to confirm cell dissociation. Add 10 mL of growth media containing 10% FBS to inactivate trypsin.</li>
	<li>Collect the cell suspension in a 50 mL conical tube and centrifuge at 400 x g for 5 min at room temperature. Gently resuspend cells in the complete cell culture medium and determine the cell count and viability using Trypan Blue (0.4%) staining. 
	NOTE: The recommended minimum cell viability for this experiment is &#62;90%.</li>
	<li>Place sterile glass coverslips in wells of a 24-well plate using autoclaved forceps. Seed cells onto coverslips at a density of 1 x 106 cells/mL and incubate the plate overnight in a humidified incubator at 37 &#176;C and 5% CO2.</li>
	<li>Change media of each well with fresh 500 &#181;L complete media before treatment. Pretreat macrophages with vehicle (DMSO or other solvents used to dissolve EIPA) or the macropinocytosis inhibitor 5-(N-ethyl-N-isopropyl)-Amiloride (EIPA, 25 &#181;M21) for 30 min.</li>
	<li>Treat cells with vehicle and stimulators of macropinocytosis to promote membrane ruffling: phorbol 12-myristate 13-acetate (PMA, 1 &#181;M, 30 min21) and macrophage colony-stimulating factor (M-CSF, 100 ng/mL, 30 min3). 
	NOTE: Alternative inhibitors [e.g., latrunculin A (1 &#181;M, 30 min) or cytochalasin D (1 &#181;M, 30 min)] and stimulators [e.g., epidermal growth factor (EGF, 100 ng/mL, 10 min) or platelet-derived growth factor (PDGF, 100 ng/mL, 15 min)] of macropinocytosis can be also used to characterize macropinocytosis in macrophages and other cell types16,27,28,29. Cells may require overnight serum starvation prior to the treatment with physiological stimulators of macropinocytosis. It is important to note that the most effective concentrations and incubation times will have to be determined to stimulate macropinocytosis in other cell types.</li>
</ol>
2. SEM fixation
<ol>
	<li>After the treatment, aspirate the media from wells and wash coverslips with ice-cold PBS twice.</li>
	<li>Fix cells (4% paraformaldehyde and 2% glutaraldehyde in 0.1 M sodium cacodylate) for 30 min at room temperature, followed by overnight incubation in the fixative at 4 &#176;C.</li>
	<li>Gently wash and incubate coverslips with 500 &#181;L of following reagents without disturbing the cell monolayer.
	<ol>
		<li>Wash once in 0.1 M sodium cacodylate and incubate for 15 min.</li>
		<li>Wash twice with dH2O with 10 min incubation in each wash.</li>
		<li>Wash twice with 25% ethanol with 10 min incubation in each wash.</li>
		<li>Wash twice with 50% ethanol with 10 min incubation in each wash.</li>
		<li>Wash twice with 75% ethanol with 10 min incubation in each wash.</li>
		<li>Wash twice with 80% ethanol with 10 min incubation in each wash.</li>
		<li>Wash twice with 95% ethanol with 10 min incubation in each wash.</li>
		<li>Wash twice with 100% ethanol. Perform 10 min of incubation in each wash. 
		NOTE: Changing/replacing of reagents should be done fast to prevent air drying of cells. Coverslips can be left in 100% ethanol for several days at 4 &#176;C. For long-term storage, add additional 100% ethanol and cover wells with parafilm to prevent air drying of the sample.</li>
	</ol>
	</li>
</ol>
3.&#160;Critical point drying
<ol>
	<li>Place coverslips in a Critical Point Dryer and cover with 100% ethanol. Press the Power button and open the CO2 tank.</li>
	<li>Press the Cool button for approximately 30 s until the temperature decreases to 0 &#176;C. Press the Fill button until a bubble appears in the chamber window.</li>
	<li>Press the Purge button until the smell of ethanol from the purge exhaust disappears. Press the Cool button again until the temperature decreases to 0 &#176;C.</li>
	<li>Repress the Fill and Purge buttons to turn them off and close the CO2 tank. Repress the Cool button to turn it off and press the Heat button.</li>
	<li>Set the temperature at 42 &#176;C and pressure at 1,200 psi. Once the pressure and temperature stabilize, press the Bleed button to allow the pressure to decrease slowly.</li>
	<li>Once the chamber pressure reaches 150 psi, press the Vent button and wait until the pressure decreases to 0 psi. Turn off the critical point dryer and remove the coverslips.</li>
	<li>Mount coverslips on SEM aluminum specimen mounts using Carbon Adhesive tabs and subject to sputter coating using gold/palladium in a sputter coater.</li>
	<li>Turn on the Power button and wait until the vacuum reaches 30 mTorr. Flush the chamber to remove humidity and air by turning off the gas switch and turning the Fine gas valve counterclockwise.</li>
	<li>Once the vacuum increases to 200 mTorr turn off the gas switch and wait until the vacuum reaches 30 mTorr. Repeat this step 3 times.</li>
	<li>Push the Timer button and adjust the Voltage knob until the gauge reads 10 mA. Remove coated coverslips from the chamber. 
	NOTE: Follow manufacturer&#39;s instruction to perform critical point drying and sputter coating of the sample.</li>
</ol>
4. Imaging and quantification
<ol>
	<li>Insert sample coverslips into the chamber of a scanning electron microscope. Close the door and press the Evac button.</li>
	<li>Open the SEM operation software. Set the accelerating voltage to 15 kv and the working distance to 10 mm.</li>
	<li>Press the Coordinates button and move around the controller until the cells appear in the center of the observation screen. Set the magnification to 3,500x and image the sample by clicking the Photo button. 
	NOTE: Use a higher level of magnification (8,500x to 16,000x) to show the plasma membrane at greater details.</li>
	<li>Take images from at least 10 random locations on the coverslips, making sure that each microscopic field contains multiple cells. Save images as .tif files.</li>
	<li>From the images, locate membrane ruffles, defined as protruding blanket-like folds of the plasma membrane, with a size greater than 500 nm (Figure 1). Count the number of ruffles per cell. 
	NOTE: C-shaped ruffles were identified as curved membrane protrusions that did not fuse on their lateral ends [(Figure 1 (M-CSF treatment) and Figure 2B]. Fused circular membrane ruffles, prior to their closure, were identified as macropinocytotic cups (Figure 2C).</li>
	<li>Normalize the number of membrane ruffles to the total number of cells in the microscopic field evaluated. Repeat this analysis for the samples from each group.</li>
</ol>

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The authors declare no conflicts of interest.

The present SEM imaging protocol provides a tool to visualize and quantify membrane ruffle formation, circular protrusions, and macropinocytic cups on the cell surface in vitro. Although the current protocol focuses on macrophages, studies have shown that membrane ruffle formation also occurs on various other cell types including dendritic cells, fibroblasts, neurons, and cancer cells11,12,14,21,30. Given this, optimization of cell culture conditions and the use of different stimulators or treatment conditions may be required for the visualization of membrane ruffles in other cell types. 
Although some leeway exists for sample preparation, the dehydration and critical point drying steps must be followed exactly to preserve the cell surface architecture. In addition, the presence of water or other solvents such as ethanol would disturb the vacuum, which is essential for SEM imaging and therefore, it must be removed in a controlled manner to keep the specimen morphology intact14.
Electron microscopes use a beam of accelerated electrons as a source of illumination. The first electron microscopes were developed when the wavelength became the limiting factor in light microscopes31. Electrons have much shorter wavelengths thus electron microscopes have a higher resolving power than light microscopes and can be used to provide high-resolution images and investigate the ultrastructure of a wide range of biological specimens. A major limitation of this method revolves around the fact that SEM provides only a snapshot of the plasma membrane morphology, as opposed to the fluorescent live cell imaging, which would allow visualization of membrane ruffles, their transition to macropinocytic cups, and formation of macropinosomes. Live cell imaging, transmission electron microscopy (TEM), and fluorescent dextran uptake assays can be used to investigate the internalization of fluorescent solutes via macropinocytosis, macropinosome translocation and maturation, and intracellular signaling mechanisms regulating macropinocytosis. Because SEM requires fixation onto coverslips, only the dorsal side and the peripheral region of the cell can be visualized, which is another limitation to this technique. 
We would like to add that sometimes it is difficult to distinguish between macropinocytosis-independent formation of membrane protrusions and macropinocytic membrane ruffling. We standardized our quantification by identifying membrane ruffles with at least 0.5 &#181;m minimum distance between any two points at the edge of the protrusion. The number of membrane ruffles per cell is highly variable and depends on numerous factors including the stimulant, its concentration, incubation time and cell type32. With PMA and M-CSF, we did see cells with multiple membrane ruffles but chose representative images that better represent our quantified data (1-2 ruffles per cell). SEM remains the gold standard technique for assessing topographical and morphological characteristics of the plasma membrane in vitro. The combination of this method along with live cell imaging and flow cytometry analysis of solute internalization provides a reliable strategy for analyzing macropinocytosis in various cell types.

Macropinocytosis is an endocytic process responsible for internalizing a large amount of extracellular fluid and its content via the formation of dynamic and actin-driven plasma membrane protrusions called membrane ruffles1. Many of these membrane ruffles form cups that close and fuse back onto the cell and separate from the plasma membrane as large, heterogeneous intracellular endosomes also known as macropinosomes1. Although macropinocytosis is induced by growth factors such as macrophage colony-stimulating factor (M-CSF) and epidermal growth factor (EGF) in a wide range of cell types, an additional unique, calcium-dependent process known as constitutive macropinocytosis has been also observed in innate immune cells2,3,4,5,6,7,8.
The ability of cells to internalize extracellular material via macropinocytosis has been shown to play an important role in a variety of physiological processes ranging from nutrient uptake to pathogen capture and antigen presentation9,10,11. However, because this process is non-selective and inducible, it has also been implicated in a number of pathological conditions. Indeed, previous studies suggested that macropinocytosis plays an important role in Alzheimer&#39;s disease, Parkinson&#39;s disease, cancer, nephrolithiasis and atherosclerosis12,13,14,15,16. Moreover, certain bacteria and viruses have shown to utilize macropinocytosis to gain entry into host cells and induce infection17,18. Interestingly, stimulation of macropinocytosis can be also exploited for targeted delivery of therapeutic agents in various disease conditions19,20.
Previous studies have explored macropinocytosis by quantifying internalized fluorescently-tagged fluid-phase markers in the absence and presence of pharmacological agents that inhibit macropinocytosis using flow cytometry and confocal imaging21,22. Currently available pharmacological tools that inhibit macropinocytosis are limited to and comprise of 1) actin polymerization inhibitors (cytochalasin D and latrunculins), 2) PI3K blockers (LY-290042 and wortmannin) and 3) inhibitors of sodium hydrogen exchangers (NHE) (amiloride and EIPA)5,14,15,23,24,25. However, because these inhibitors have endocytosis independent effects, it is difficult to selectively determine the contribution of macropinocytosis to solute uptake and disease pathogenesis especially in vivo21.
Scanning electron microscopy (SEM) is a type of electron microscope that produces ultra-high-resolution images of cells using a focused beam of electrons26. In macropinocytosis research, SEM imaging is regarded as the gold standard technique to visualize topographical and morphological characteristics of the plasma membrane, quantify membrane ruffle formation, and investigate their progression towards macropinosome internalization. Furthermore, scanning electron microscopy combined with the quantification of solute uptake, in the presence and absence of macropinocytosis blockers, provides a reliable strategy to examine macropinocytotic solute internalization in vitro. This paper provides a detailed protocol on how to prepare cells for SEM, visualize the cell surface, quantify ruffle formation, and examine their progress towards cup closure and macropinosome internalization.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.5% Trypsin-EDTA</td>
			<td>Gibco</td>
			<td>15400-054</td>
			<td></td>
		</tr>
		<tr>
			<td>2% Glutaraldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>16320</td>
			<td></td>
		</tr>
		<tr>
			<td>4% Paraformaldehyde</td>
			<td>Santa Cruz Biotechnology</td>
			<td>281692</td>
			<td></td>
		</tr>
		<tr>
			<td>5-(N-ethyl-N-isopropyl)-Amiloride</td>
			<td>Sigma Life Science</td>
			<td>A3085</td>
			<td></td>
		</tr>
		<tr>
			<td colspan="2">Accuri C6 Flow Cytometer</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Carbon Adhesive Tabs</td>
			<td>Electron Microscopy Sciences</td>
			<td>77825-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Dimethyl Sulfoxide</td>
			<td>Corning</td>
			<td>25-950-CQC</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle Medium</td>
			<td>Cytiva Life Sciences</td>
			<td>SH30022.01</td>
			<td></td>
		</tr>
		<tr>
			<td>Falcon 24-well Clear Flat Bottom TC-treated Multiwell Cell Culture Plate</td>
			<td>Falcon</td>
			<td>353047</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>Gemini Bio</td>
			<td>900-108</td>
			<td></td>
		</tr>
		<tr>
			<td>FitC-dextran</td>
			<td>Thermo Fisher Scientific</td>
			<td>D1823</td>
			<td></td>
		</tr>
		<tr>
			<td>FM 4-64</td>
			<td>Thermo Fisher Scientific</td>
			<td>T13320</td>
			<td></td>
		</tr>
		<tr>
			<td>HERAcell 150i CO2 incubator</td>
			<td>Thermo Fisher Scientific</td>
			<td>51026282</td>
			<td></td>
		</tr>
		<tr>
			<td>Hummer Model 6.2 Sputter Coater</td>
			<td>Anatech USA</td>
			<td>58565</td>
			<td></td>
		</tr>
		<tr>
			<td colspan="2">JSM-IT500HR scanning electron microscope</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Cover Glass</td>
			<td>Thermo Fisher Scientific</td>
			<td>12-545-82</td>
			<td></td>
		</tr>
		<tr>
			<td>Pen Strep</td>
			<td>Gibco</td>
			<td>15140-122</td>
			<td></td>
		</tr>
		<tr>
			<td>phorbol 12-myristate 13-acetate</td>
			<td>Millipore Sigma</td>
			<td>524400</td>
			<td></td>
		</tr>
		<tr>
			<td>RAW 264.7 macrophage</td>
			<td>ATCC</td>
			<td>ATCC TIB-71</td>
			<td></td>
		</tr>
		<tr>
			<td>Recombinant Human M-CSF</td>
			<td>Peprotech</td>
			<td>300-25</td>
			<td></td>
		</tr>
		<tr>
			<td>Samdri-790 Critical Point Dryer</td>
			<td>Tousimis Research Corporation</td>
			<td>8778B</td>
			<td></td>
		</tr>
		<tr>
			<td>SEM Aluminum Specimen Mounts</td>
			<td>Electron Microscopy Sciences</td>
			<td>75220</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Cacodylate</td>
			<td>Electron Microscopy Sciences</td>
			<td>12300</td>
			<td></td>
		</tr>
		<tr>
			<td>Texas red-dextra</td>
			<td>Thermo Fisher Scientific</td>
			<td>D1864</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypan Blue Solution</td>
			<td>Thermo Fisher Scientific</td>
			<td>15250061</td>
			<td></td>
		</tr>
		<tr>
			<td>Zeiss LSM 780 confocal microscope</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

visualizing membrane ruffle formation using scanning electron microscopy

Membrane ruffling is the formation of motile plasma membrane protrusions containing a meshwork of newly polymerized actin filaments. Membrane ruffles may form spontaneously or in response to growth factors, inflammatory cytokines, and phorbol esters. Some of the membrane protrusions may reorganize into circular membrane ruffles that fuse at their distal margins and form cups that close and separate into the cytoplasm as large, heterogeneous vacuoles called macropinosomes. During the process, ruffles trap extracellular fluid and solutes that internalize within macropinosomes. High-resolution scanning electron microscopy (SEM) is a commonly used imaging technique to visualize and quantify membrane ruffle formation, circular protrusions, and closed macropinocytic cups on the cell surface. The following protocol describes the cell culture conditions, stimulation of the membrane ruffle formation in vitro, and how to fix, dehydrate, and prepare cells for imaging using SEM. Quantification of membrane ruffling, data normalization, and stimulators and inhibitors of membrane ruffle formation are also described. This method can help answer key questions about the role of macropinocytosis in physiological and pathological processes, investigate new targets that regulate membrane ruffle formation, and identify yet uncharacterized physiological stimulators as well as novel pharmacological inhibitors of macropinocytosis.

Here, we describe the results from the presented technique. Representative SEM images shown in Figure 1 demonstrate membrane ruffle formation in RAW 264.7 macrophages following treatment with PMA and M-CSF. Images were first captured at a magnification of 3,500x for quantification purposes and then at higher levels of magnification (8,500x to 16,000x) to show the plasma membrane at greater details. Pretreatment of macrophages with the macropinocytosis inhibitor EIPA attenuated membrane ruffle formation (Figure 1). Macropinocytosis-associated plasma membrane activities can be divided into five consecutive steps: 1) initiation of membrane ruffling, 2) circularization, 3) cup formation, 4) cup closure, and 5) internalization of extracellular fluid and its associated solutes via macropinosome formation. Figure 2 shows representative images of these morphologically distinct plasma membrane activities following M-CSF stimulation. Large sheet-like ruffles (Figure 2A), circularized C-shaped ruffles (Figure 2B), and macropinocytic cups (Figure 2C) were captured at a magnification of 6,000x to 7,000x. Scanning electron microscopy can be used to provide high-resolution images of the cell surface but cannot be utilized to visualize internalized macropinosomes. Quantification of membrane ruffles following PMA and M-CSF treatment is shown in Figure 3. Macropinosome formation can be confirmed by alternative imaging techniques, including confocal microscopy, as shown in Figure 4. Finally, SEM quantification of membrane ruffling should be complemented by flow cytometry quantification of a fluorescent fluid-phase marker (e.g., FITC- or Texas red-dextran) to confirm stimulation of macropinocytosis (Figure 5).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62658/62658fig01.jpg" /> 
Figure 1: Scanning electron microscopy images of RAW 264.7 macrophages. RAW 264.7 macrophages were pre-treated with vehicle (DMSO control) or EIPA (25 &#181;M) for 30 min and incubated with PMA (1 &#181;M, 30 min) or M-CSF (100 ng/mL, 30 min) to stimulate membrane ruffling. Images on the right show higher magnification of the cell surface. Red arrows: membrane ruffles. Green arrow: c-shaped ruffle. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62658/62658fig01large.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/62658/62658fig02.jpg" /> 
Figure 2: Morphological stages of macropinocytosis captured by scanning electron microscopy. RAW 264.7 macrophages were treated with M-CSF (100 ng/mL) for 30 min and cells were processed for SEM imaging. (A) sheet-like membrane protrusion, (B) C-shaped membrane ruffle, and (C) macropinocytic cup. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62658/62658fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62658/62658fig03.jpg" /> 
Figure 3: Quantification of membrane ruffle formation. RAW 264.7 macrophages were pre-treated with vehicle (DMSO control) or EIPA (25 &#181;M) for 30 min and incubated with PMA (1 &#181;M, 30 min) or M-CSF (100 ng/mL, 30 min) to stimulate membrane ruffling. Bar graph shows the number of membrane ruffles normalized to total cell number (n = 3). Data represent the mean &#177; SEM. One-way ANOVA; *p &#60; 0.05 vs. vehicle, #p &#60; 0.05 vs. PMA or M-CSF treatment. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62658/62658fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62658/62658fig04.jpg" /> 
Figure 4: Macropinosome formation. Cells were pre-treated with PMA for 30 min and incubated with Texas red-dextran (25 &#181;g/mL, red) and FM4-64 (5 &#181;g/mL, green) for 10 min. Imaging were performed using a confocal microscope. Blue arrows: membrane ruffling, yellow arrows: macropinosome containing Texas red-dextran, green arrows: macropinosomes. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62658/62658fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62658/62658fig05.jpg" /> 
Figure 5: Quantification of macropinocytosis. RAW 264.7 macrophages were pre-treated with vehicle (DMSO control) or EIPA (25 &#181;M) for 30 min and incubated with PMA (1 &#181;M) and FITC-dextran (100 &#181;g/mL; 70,000 MW) for 2 h. Bar graph shows the mean fluorescence intensity (MFI) fold change normalized to vehicle treatment (n = 3, performed in triplicates). Data represent the mean &#177; SEM. One-way ANOVA; *p &#60; 0.05 vs. vehicle, #p &#60; 0.05 vs. PMA. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62658/62658fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Visualizing Membrane Ruffle Formation using Scanning Electron Microscopy at JoVE.com

Visualizing Membrane Ruffle Formation using Scanning Electron Microscopy

Macropinocytosis is a highly conserved endocytic process initiated by the formation of F-actin-rich sheet-like membrane projections, also known as membrane ruffles. Increased rate of macropinocytotic solute internalization has been implicated in various pathological conditions. This protocol presents a method to quantify membrane ruffle formation in vitro using scanning electron microscopy.

Membrane ruffling is the formation of motile plasma membrane protrusions containing a meshwork of newly polymerized actin filaments. Membrane ruffles may ...

visualizing-membrane-ruffle-formation-using-scanning-electron

Nanyang Technological University

Research

JoVE Journal

Medicine

2.4K Views.  Medical College of Georgia at Augusta University. Macropinocytosis is a highly conserved endocytic process initiated by the formation of F-actin-rich sheet-like membrane projections, also known as membrane ruffles. Increased rate of macropinocytotic solute internalization has been implicated in various pathological conditions. This protocol presents a method to quantify membrane ruffle formation in vitro using scanning electron microscopy. 

Video: Visualizing Membrane Ruffle Formation using Scanning Electron Microscopy

Oral Biofilm Analysis of Palatal Expanders by Fluorescence In-Situ Hybridization and Confocal Laser Scanning Microscopy

Confocal laser scanning microscopy (CLSM) of natural heterogeneous biofilm is today facilitated by a comprehensive range of staining techniques, one of them being fluorescence in situ hybridization (FISH).1,2 We performed a pilot study in which oral biofilm samples collected from fixed orthodontic appliances (palatal expanders) were stained by FISH, the objective being to assess the three-dimensional organization of natural biofilm and plaque accumulation.3,4 FISH creates an opportunity to stain cells in their native biofilm environment by the use of fluorescently labeled 16S rRNA-targeting probes.4-7,19 Compared to alternative techniques like immunofluorescent labeling, this is an inexpensive, precise and straightforward labeling technique to investigate different bacterial groups in mixed biofilm consortia.18,20 General probes were used that bind to Eubacteria (EUB338 + EUB338II + EUB338III; hereafter EUBmix),8-10 Firmicutes (LGC354 A-C; hereafter LGCmix),9,10 and Bacteroidetes (Bac303).11 In addition, specific probes binding to Streptococcus mutans (MUT590)12,13 and Porphyromonas gingivalis (POGI)13,14 were used. The extreme hardness of the surface materials involved (stainless steel and acrylic resin) compelled us to find new ways of preparing the biofilm. As these surface materials could not be readily cut with a cryotome, various sampling methods were explored to obtain intact oral biofilm. The most workable of these approaches is presented in this communication. Small flakes of the biofilm-carrying acrylic resin were scraped off with a sterile scalpel, taking care not to damage the biofilm structure. Forceps were used to collect biofilm from the steel surfaces. Once collected, the samples were fixed and placed directly on polysine coated glass slides. FISH was performed directly on these slides with the probes mentioned above. Various FISH protocols were combined and modified to create a new protocol that was easy to handle.5,10,14,15 Subsequently the samples were analyzed by confocal laser scanning microscopy. Well-known configurations3,4,16,17 could be visualized, including mushroom-style formations and clusters of coccoid bacteria pervaded by channels. In addition, the bacterial composition of these typical biofilm structures were analyzed and 2D and 3D images created.

We present a protocol for structural and compositional analysis of natural oral biofilm from orthodontic appliances with in situ hybridization (FISH) and confocal laser scanning microscopy (CLSM). Oral biofilm samples were collected from palatal expanders, scraping acrylic-resin flakes off their surface and referring them for molecular processing.

Confocal laser scanning microscopy (CLSM) of natural heterogeneous biofilm is today facilitated by a comprehensive range of staining techniques, one of ...

Formation of Human Prostate Epithelium Using Tissue Recombination of Rodent Urogenital Sinus Mesenchyme and Human Stem Cells

Progress in prostate cancer research is severely limited by the availability of human-derived and hormone-na&iuml;ve model systems, which limit our ability to understand genetic and molecular events underlying prostate disease initiation. Toward developing better model systems for studying human prostate carcinogenesis, we and others have taken advantage of the unique pro-prostatic inductive potential of embryonic rodent prostate stroma, termed urogenital sinus mesenchyme (UGSM). When recombined with certain pluripotent cell populations such as embryonic stem cells, UGSM induces the formation of normal human prostate epithelia in a testosterone-dependent manner. Such a human model system can be used to investigate and experimentally test the ability of candidate prostate cancer susceptibility genes at an accelerated pace compared to typical rodent transgenic studies. Since Human embryonic stem cells (hESCs) can be genetically modified in culture using inducible gene expression or siRNA knock-down vectors prior to tissue recombination, such a model facilitates testing the functional consequences of genes, or combinations of genes, which are thought to promote or prevent carcinogenesis.
The technique of isolating pure populations of UGSM cells, however, is challenging and learning often requires someone with previous expertise to personally teach. Moreover, inoculation of cell mixtures under the renal capsule of an immunocompromised host can be technically challenging. Here we outline and illustrate proper isolation of UGSM from rodent embryos and renal capsule implantation of tissue mixtures to form human prostate epithelium. Such an approach, at its current stage, requires in vivo xenografting of embryonic stem cells; future applications could potentially include in vitro gland formation or the use of induced pluripotent stem cell populations (iPSCs).

To unravel the earliest molecular mechanisms underlying prostate cancer initiation, novel and innovative human model systems and approaches are desperately needed. The potential of pre-prostatic urogenital sinus mesenchyme (UGSM) to induce pluripotent stem cell populations to form human prostate epithelium is a powerful experimental tool in prostate research.

Progress in prostate cancer research is severely  limited by the availability of human-derived and hormone-na&iuml;ve model systems,  which limit our ...

Using a Laminating Technique to Perform Confocal Microscopy of the Human Sclera

The sclera is a dense connective tissue that covers and protects the eye. It mainly consists of dense collagen bundles (types I, III, IV, V, VI, and VII). Due to its autofluorescence, opaqueness, and thickness, it has not been found suitable for confocal microscopy. An alternative approach to the one presented here, which uses formalin-fixed sclera embedded in paraffin for immunohistochemistry, has technical challenges, especially when preheating the tissue for antigen retrieval. Since the sclera is relatively poor in both cells and vessels, the use of larger tissue samples was explored to help prevent overlooking cells and to understand their localization in relation to vessels and other anatomical sites. To allow for the analysis of larger tissue samples under the confocal microscope, a laminating technique was performed to create thin layers from the sclera. Following the analysis of results of CD31 blood vessels and lymphatic vessel endothelial hyaluronan receptor 1 (LYVE1) positive cells, for which approval for scientific examination was obtained, the advantages and limitations of this method are discussed.

Human sclera tissue is mainly collagen; therefore, it is not easily usable for immunohistochemistry. To achieve the goal of performing immunohistochemistry for confocal microscopy of scleral tissue, a laminating technique was used.


The sclera is a dense connective tissue that covers and protects the eye. It mainly consists of dense collagen bundles (types I, III, IV, V, VI, and ...

Quantitative Visualization of Leukocyte Infiltrate in a Murine Model of Fulminant Myocarditis by Light Sheet Microscopy

Light-sheet fluorescence microscopy (LSFM), in combination with chemical clearing protocols, has become the gold standard for analyzing fluorescently labelled structures in large biological specimens, and is down to cellular resolution. Meanwhile, the constant refinement of underlying protocols and the enhanced availability of specialized commercial systems enable us to investigate the microstructure of whole mouse organs and even allow for the characterization of cellular behavior in various live-cell imaging approaches. Here, we describe a protocol for the spatial whole-mount visualization and quantification of the CD45+ leukocyte population in inflamed mouse hearts. The method employs a transgenic mouse strain (CD11c.DTR)that has recently been shown to serve as a robust, inducible model for the study of the development of fulminant fatal myocarditis, characterized by lethal cardiac arrhythmias. This protocol includes myocarditis induction, intravital antibody-mediated cell staining, organ preparation, and LSFM with subsequent computer-assisted image post-processing. Although presented as a highly-adapted method for our particular scientific question, the protocol represents the blueprint of an easily adjustable system that can also target completely different fluorescent structures in other organs and even in other species.

Here, we describe a light-sheet microscopy approach to visualize the cardiac CD45+ leukocyte infiltrate in a murine model of aseptic fulminant myocarditis, which is induced by the intratracheal diphtheria toxin treatment of CD11c.DTR mice.

Light-sheet fluorescence microscopy (LSFM), in combination with chemical clearing protocols, has become the gold standard for analyzing fluorescently ...

Preparation and In Vitro Characterization of Magnetized miR-modified Endothelial Cells

To date, the available surgical and pharmacological treatments for cardiovascular diseases (CVD) are limited and often palliative. At the same time, gene and cell therapies are highly promising alternative approaches for CVD treatment. However, the broad clinical application of gene therapy is greatly limited by the lack of suitable gene delivery systems. The development of appropriate gene delivery vectors can provide a solution to current challenges in cell therapy. In particular, existing drawbacks, such as limited efficiency and low cell retention in the injured organ, could be overcome by appropriate cell engineering (i.e., genetic) prior to transplantation. The presented protocol describes the efficient and safe transient modification of endothelial cells using a polyethyleneimine superparamagnetic magnetic nanoparticle (PEI/MNP)-based delivery vector. Also, the algorithm and methods for cell characterization are defined. The successful intracellular delivery of microRNA (miR) into human umbilical vein endothelial cells (HUVECs) has been achieved without affecting cell viability, functionality, or intercellular communication. Moreover, this approach was proven to cause a strong functional effect in introduced exogenous miR. Importantly, the application of this MNP-based vector ensures cell magnetization, with accompanying possibilities of magnetic targeting and non-invasive MRI tracing. This may provide a basis for magnetically guided, genetically engineered cell therapeutics that can be monitored non-invasively with MRI.

Preparation and In Vitro Characterization of Magnetized miR-modified Endothelial Cells

This manuscript describes the efficient, non-viral delivery of miR to endothelial cells by a PEI/MNP vector and their magnetization. Thus, in addition to genetic modification, this approach allows for magnetic cell guidance and MRI detectability. The technique can be used to improve the characteristics of therapeutic cell products.

To date, the available surgical and pharmacological treatments for cardiovascular diseases (CVD) are limited and often palliative. At the same time, gene ...

Scanning Skeletal Remains for Bone Mineral Density in Forensic Contexts

The purpose of this paper is to introduce a promising, novel method to aid in the assessment of bone quality in forensically relevant skeletal remains. BMD is an important component of bone&#39;s nutritional status and in skeletal remains of both juveniles and adults, and it can provide information about bone quality. For adults remains, it can provide information on pathological conditions or when bone insufficiency may have occurred. In juveniles, it provides a useful metric to elucidate cases of fatal starvation or neglect, which are generally difficult to identify. This paper provides a protocol for the anatomical orientation and analysis of skeletal remains for scanning via dual-energy X-ray absorptiometry (DXA). Three case studies are presented to illustrate when DXA scans can be informative to the forensic practitioner. The first case study presents an individual with observed longitudinal fractures in the weight bearing bones and DXA is used to assess bone insufficiency. BMD is found to be normal suggesting another etiology for the fracture pattern present. The second case study employed DXA to investigate suspected chronic malnutrition. The BMD results are consistent with results from long bone lengths and suggest the juvenile had suffered from chronic malnutrition. The final case study provides an example where fatal starvation in a fourteen-month infant is suspected, which supports autopsy findings of fatal starvation. DXA scans showed low bone mineral density for chronological age and is substantiated by traditional assessments of infant health. However, when dealing with skeletal remains taphonomic alterations should be considered before applying this method.

Bone mineral density (BMD) is an important factor in understanding nutritional intake. For human skeletal remains, it is a useful metric to assess quality of life in both juveniles and adults, particularly in fatal starvation and neglect cases. This paper provides guidelines for scanning human skeletal remains for forensic purposes.

The purpose of this paper is to introduce a promising, novel method to aid in the assessment of bone quality in forensically relevant skeletal remains. ...

Micro-dissection of Enamel Organ from Mandibular Incisor of Rats Exposed to Environmental Toxicants

Enamel defects resulting from environmental conditions and ways of life are public health concerns because of their high prevalence. These defects result from altered activity of cells responsible for enamel synthesis named ameloblasts, which present in enamel organ. During amelogenesis, ameloblasts follow a specific and precise sequence of events of proliferation, differentiation, and death. A rat continually growing incisors is a suitable experimental model to study ameloblast activity and differentiation stages in physiological and pathological conditions. Here, we describe a reliable and consistent method to micro-dissect enamel organ of rats exposed to environmental toxicants. The micro-dissected dental epithelia contain secretion- and maturation-stage ameloblasts that may be used for qualitative experiments, such as immunohistochemistry assays and in situ hybridization, as well as for quantitative analyses such as RT-qPCR, RNA-seq, and Western blotting.

Understanding enamel formation and possible alterations requires the study of ameloblast activity. Here, we describe a reliable and consistent method to micro-dissect enamel organs containing secretion- and maturation-stage ameloblasts that may be used for further quantitative and qualitative experimental procedures.

Enamel defects resulting from environmental conditions and ways of life are public health concerns because of their high prevalence. These defects result ...

Noninvasive Determination of Vortex Formation Time Using Transesophageal Echocardiography During Cardiac Surgery

Trans-mitral blood flow produces a three-dimensional rotational body of fluid, known as a vortex ring, that enhances the efficiency of left ventricular (LV) filling compared with a continuous linear jet. Vortex ring development is most often quantified with vortex formation time (VFT), a dimensionless parameter based on fluid ejection from a rigid tube. Our group is interested in factors that affect LV filling efficiency during cardiac surgery. In this report, we describe how to use standard two-dimensional (2D) and Doppler transesophageal echocardiography (TEE) to noninvasively derive the variables needed to calculate VFT. We calculate atrial filling fraction (&#946;) from velocity-time integrals of trans-mitral early LV filling and atrial systole blood flow velocity waveforms measured in the mid-esophageal four-chamber TEE view. Stroke volume (SV) is calculated as the product of the diameter of the LV outflow track measured in the mid-esophageal long axis TEE view and the velocity-time integral of blood flow through the outflow track determined in the deep transgastric view using pulse-wave Doppler. Finally, mitral valve diameter (D) is determined as the average of major and minor axis lengths measured in orthogonal mid-esophageal bicommissural and long axis imaging planes, respectively. VFT is then calculated as 4 &#215; (1-&#946;) &#215; SV/(&#960;D3). We have used this technique to analyze VFT in several groups of patients with differing cardiac abnormalities. We discuss our application of this technique and its potential limitations and also review our results to date. Noninvasive measurement of VFT using TEE is straightforward in anesthetized patients undergoing cardiac surgery. The technique may allow cardiac anesthesiologists and surgeons to assess the impact of pathological conditions and surgical interventions on LV filling efficiency in real time.

We describe a protocol to measure vortex formation time, an index of left ventricular filling efficiency, using standard transesophageal echocardiography techniques in patients undergoing cardiac surgery. We apply this technique to analyze vortex formation time in several groups of patients with differing cardiac pathologies.

Trans-mitral blood flow produces a three-dimensional rotational body of fluid, known as a vortex ring, that enhances the efficiency of left ventricular ...

Immunofluorescence Labelling of Human and Murine Neutrophil Extracellular Traps in Paraffin-Embedded Tissue

Neutrophil granulocytes, also called polymorphonuclear leukocytes (PMN) due to their lobulated nucleus, are the most abundant type of leukocytes. They mature in the bone marrow and are released into the peripheral blood, where they circulate for about 6&#8722;8 h; however, in tissue, they can survive for days. By diapedesis through the endothelium, they leave the blood stream, enter tissues, and migrate towards the site of an infection following chemotactic gradients.&#160;Neutrophils can combat invading microorganisms by phagocytosis,&#160;degranulation, and generation of neutrophil extracellular traps (NETs). This protocol will help to detect NETs in paraffin-embedded tissue. NETs are the result of a process called NETosis, which leads to the release of nuclear, granular, and cytoplasmic components either from living (vital NETosis) or dying (suicidal NETosis) neutrophils. In vitro, NETs form cloud-like structures, which occupy a space several times larger than that of the cells from which they descended. The backbone of NETs is chromatin, to which a selection of proteins and peptides originating from granules and cytoplasm are bound. Thereby, a high local concentration of toxic compounds is maintained so that NETs can capture and inactivate a variety of pathogens including bacteria, fungi, viruses, and parasites, while diffusion of the highly active NET components leading to damage in neighboring tissue is limited. Nevertheless, in recent years it has become apparent that NETs, if generated in abundance or cleared insufficiently, do have pathological potential ranging from autoimmune diseases to cancer. Thus, detection of NETs in tissue samples may have diagnostic significance, and the detection of NETs in diseased tissue can influence the treatment of patients. Since paraffin-embedded tissue samples are the standard specimen used for pathological analysis, it was sought to establish a protocol for fluorescent staining of NET components in paraffin-embedded tissue using commercially available antibodies.

Neutrophil extracellular traps (NETs) are three-dimensional structures generated by stimulated neutrophil granulocytes. It has become clear in recent years that NETs are involved in a wide variety of diseases. Detection of NETs in tissue may have diagnostic relevance, so standardized protocols for labelling NET components are required.

Neutrophil granulocytes, also called polymorphonuclear leukocytes (PMN) due to their lobulated nucleus, are the most abundant type of leukocytes. They ...

Visualizing and Quantifying Pharmaceutical Compounds within Skin using Coherent Raman Scattering Imaging

Cutaneous pharmacokinetics (cPK) after topical formulation application has been a research area of particular interest for regulatory and drug development scientists to mechanistically understand topical bioavailability (BA). Semi-invasive techniques, such as tape-stripping, dermal microdialysis, or dermal open-flow microperfusion, all quantify macroscale cPK. While these techniques have provided vast cPK knowledge, the community lacks a mechanistic understanding of active pharmaceutical ingredient (API) penetration and permeation at the cellular level. 
One noninvasive approach to address microscale cPK is coherent Raman scattering imaging (CRI), which selectively targets intrinsic molecular vibrations without the need for extrinsic labels or chemical modification. CRI has two main methods-coherent anti-Stokes Raman scattering (CARS) and stimulated Raman scattering (SRS)-that enable sensitive and selective quantification of APIs or inactive ingredients. CARS is typically utilized to derive structural skin information or visualize chemical contrast. In contrast, the SRS signal, which is linear with molecular concentration, is used to quantify APIs or inactive ingredients within skin stratifications. 
Although mouse tissue has commonly been utilized for cPK with CRI, topical BA and bioequivalence (BE) must ultimately be assessed in human tissue before regulatory approval. This paper presents a methodology to prepare and image ex vivo skin to be used in quantitative pharmacokinetic CRI studies in the evaluation of topical BA and BE. This methodology enables reliable and reproducible API quantification within human and mouse skin over time. The concentrations within lipid-rich and lipid-poor compartments, as well as total API concentration over time are quantified; these are utilized for estimates of micro- and macroscale BA and, potentially, BE.

A coherent Raman scattering imaging methodology to visualize and quantify pharmaceutical compounds within the skin is described. This paper describes skin tissue preparation (human and mouse) and topical formulation application, image acquisition to quantify spatiotemporal concentration profiles, and preliminary pharmacokinetic analysis to assess topical drug delivery.

Cutaneous pharmacokinetics (cPK) after topical formulation application has been a research area of particular interest for regulatory and drug development ...

Visualizing Membrane Ruffle Formation using Scanning Electron Microscopy

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Chapters in this video

Oral Biofilm Analysis of Palatal Expanders by Fluorescence In-Situ Hybridization and Confocal Laser Scanning Microscopy

Formation of Human Prostate Epithelium Using Tissue Recombination of Rodent Urogenital Sinus Mesenchyme and Human Stem Cells

Using a Laminating Technique to Perform Confocal Microscopy of the Human Sclera

Quantitative Visualization of Leukocyte Infiltrate in a Murine Model of Fulminant Myocarditis by Light Sheet Microscopy

Preparation and In Vitro Characterization of Magnetized miR-modified Endothelial Cells

Scanning Skeletal Remains for Bone Mineral Density in Forensic Contexts

Micro-dissection of Enamel Organ from Mandibular Incisor of Rats Exposed to Environmental Toxicants

Noninvasive Determination of Vortex Formation Time Using Transesophageal Echocardiography During Cardiac Surgery

Immunofluorescence Labelling of Human and Murine Neutrophil Extracellular Traps in Paraffin-Embedded Tissue

Visualizing and Quantifying Pharmaceutical Compounds within Skin using Coherent Raman Scattering Imaging