Name: quantification of monocyte transmigration and foam cell formation from individuals with chronic inflammatory conditions
Uploaded: 2017-10-17T12:30:00.000+00:00
Duration: 9 min 41 s
Description: Watch this Scientific Journal Video about Quantification of Monocyte Transmigration and Foam Cell Formation from Individuals with Chronic Inflammatory Conditions at JoVE.com

The authors gratefully acknowledge the work of Prof. William Muller and Dr. Clare Westhorpe for their key role in development of earlier iterations of this model. The authors would also like to thank the AMREP Flow Cytometry core for the sorting of monocyte subsets and the Alfred Hospital Infectious Disease Unit clinical research nurses for the recruitment of HIV+ individuals for some studies. The authors gratefully acknowledge the contribution to this work of the Victoria Operational Infrastructure Support Program received by the Burnet Institute. TAA is supported by an RMIT University Vice-Chancellor&#39;s Postdoctoral Fellowship. This work was supported by NHMRC project grant 1108792 awarded to AJ and AH. TK is supported by NIH grants NIH K08AI08272, NIH/NCATS Grant # UL1TR000124.

NOTE: All experiments using human biological samples were performed with ethics approval from the Alfred Hospital Human Ethics Committee, Melbourne. All experiments were performed in Class II Biosafety cabinets unless specified. &#34;Prewarmed&#34; refers to reagents warmed to 37 &#176;C in a waterbath.
1. Preparation of Type I Fibrous Collagen Gels: Day 1
<ol>
	<li>Prepare polymerized collagen gels by sequentially adding and mixing 35.7 mM NaOH, 0.71 x M199, 4.58 mM acetic acid and 1.71 mg/mL type I fibrous collagen into a 5 mL polystyrene tube as per Table 1. 
	NOTE: Ensure that the collagen is well-mixed by gently pipetting up and down 5 times in order to stop the formation of collagen &#39;pockets&#39; in the gel mixture. Prepare 4-6 gels per test condition for microscopy or 15 gels per test condition for flow cytometry.</li>
	<li>Once well mixed, aliquot 50 &#181;L of collagen gel mixture into each well of a sterile flat-bottomed 96-well tissue culture plate. Do not use the outside rows and columns, but fill these with 200 &#181;L of 1x Dubecco&#39;s phosphate buffered saline (PBS) to protect the gels from desiccation. Incubate plates at 37 &#176;C/5% CO2 for 2 h to allow the collagen to polymerize. 
	Note: Place the plates directly on clean metal racks in the incubator to ensure even heat distribution.</li>
	<li>Following incubation, overlay the gels with 150 &#181;L of 1x supplemented M199 (see Table of Materials) and incubate for 5 days until use.</li>
</ol>
2. Expansion of Stored HUVEC: Day 1
<ol>
	<li>Label and coat a 10 cm diameter Petri dish with 1 mL of 50 &#181;g/mL fibronectin diluted in 1x PBS and incubate at room temperature for 10 min.</li>
	<li>Thaw aliquots of cryopreserved primary human umbilical vein endothelial cells (HUVEC, 1.0 x 106 cells) in 10 mL M20. 
	NOTE: HUVEC should be prepared as previously described21 and used at a low passage number (&#60; passage 4). HUVEC may be replaced with human coronary artery endothelial cells here.</li>
	<li>Resuspend HUVEC in 10 mL M20 and add to fibronectin coated dish, aspirating excess fibronectin prior to addition of cells, and culture to confluence (approximately 5 days), replacing the media on Day 3.</li>
</ol>
3. Culturing HUVEC Monolayer on Collagen Gels: Day 5
<ol>
	<li>On Day 5, detach HUVEC by aspirating culture supernatant from the Petri dish and washing away serum-containing media with 10 mL of serum-free M199.</li>
	<li>Add 5 mL 0.05% trypsin/0.53 EDTA in M199 to HUVEC and incubate for 1-2 min at room temperature, shaking gently until the cells detach.</li>
	<li>Once detached, quickly rinse Petri dish with 5 mL M20 and transfer the media containing cells to a 10 mL tube.</li>
	<li>Centrifuge samples at 300 x g for 5 min at room temperature, aspirate supernatant, resuspend cells in 200 &#181;L of M20, and count cells using a hemocytometer.</li>
	<li>Resuspend the cells to 2.0 x 105 cells/mL in M20, aspirate the M199 on the gels from the 96-well plate (step 1.3) and add 100 &#181;L of resuspended HUVEC (2.0 x 104 cells) to each collagen gel prepared above (step 1.2). Culture plates for a further 3 days at 37 &#176;C/5% CO2. 
	NOTE: The integrity of the HUVEC monolayer may be confirmed after 3 days by silver nitrate staining22 or immunohistochemistry for tight junction proteins23 at this stage. Additional gels must be prepared for this purpose.</li>
</ol>
4. Activation of HUVEC Monolayer and Isolation/Activation of Monocytes for Transmigration: Day 8
<ol>
	<li>On Day 8, activate each HUVEC monolayer prior to the addition of monocytes by aspirating the M20 media overlaying the gels and adding 100 &#181;L of 10 ng/mL human TNF&#945; in M20 per gel. Incubate at 37 &#176;C/5% CO2 for 4 h. 
	NOTE: Non-activated HUVEC conditions may also be included if required as controls.</li>
	<li>During the 4 h HUVEC activation step, isolate monocytes from PBMCs using magnetic bead techniques to negatively select for cells as per the manufacturer&#39;s instructions. A minimum of 6.5 x 106 PBMCs should be used at this step in order to reliably recover 3.0 x 105 monocytes required for one condition, as monocytes typically account for approximately 10-15% of PBMCs. 
	NOTE: To evaluate the effect of monocyte activation prior to monocyte transmigration and foam cell formation, monocytes can be activated at this stage. Monocytes can be isolated from thawed PBMCs if required (i.e., stored patient samples). Monocyte subsets isolated by FACS sorting can be prepared for addition to gels (Figure 1).</li>
</ol>
5. Transmigration of Primary Human Monocytes: Day 8
<ol>
	<li>To measure monocyte transmigration, remove TNF&#945;-containing media and wash gels twice with 100 &#181;L M199 by adding and removing media. Following washing, add 2.0 x 105 freshly isolated or thawed and washed cryopreserved PBMCs or 5.0 x 104 freshly isolated monocytes or purified monocyte subsets to HUVEC monolayers in 100 &#181;L M20. Incubate for 1 h at 37 &#176;C and 5% CO2 to allow the forward transmigration of monocytes into the gel. It is best to allow 6 wells for every experimental condition examined. 
	NOTE: To evaluate the effect of autologous serum on transmigration and foam cell formation, incubate cells with M20 containing the desired concentration of heat-inactivated donor serum and compare to conditions with a pooled human serum control. Leukocytes other than monocytes may be added at this stage. If investigating the impact of specific lipids/lipid species (e.g., HDL, oxHDL), perform all following steps with serum-free media containing 20-50 &#181;g/mL lipids.</li>
	<li>After 1 h of forward transmigration, collect the non-transmigrated cells by washing gels twice with 100 &#181;L of prewarmed 1 mM EGTA in 1x PBS, and once with 100 &#181;L M199 (same centrifuge conditions), pooling the supernatants from each well of the same experimental condition (usually 6 wells) into a 1.5 mL microcentrifuge tube on ice. Centrifuge the non-transmigrated cells at 4 &#176;C, 300 x g for 5 min and resuspend in 30 &#181;L 1x PBS.</li>
	<li>Count the cells collected in the supernatant to determine the number of forward transmigrated cells.</li>
	<li>The percentage of forward transmigrated cells is determined as follows: 
	<img alt="Equation 1" src="/files/ftp_upload/56293/56293eq1.jpg" /> <img alt="Equation 2" src="/files/ftp_upload/56293/56293eq2.jpg" /> <img alt="Equation 3" src="/files/ftp_upload/56293/56293eq3.jpg" /></li>
	<li>Cover the washed gels containing transmigrated cells with 100 &#181;L M20 and incubate for a further 48 h.</li>
	<li>After 48 h, collect the supernatant and wash non-transmigrated cells twice with 100 &#181;L prewarmed 1 mM EGTA/PBS, collecting and pooling the supernatant from each condition as in step 5.2. 
	NOTE: The phenotype of reverse transmigrated cells may be tested on these cells following standard flow cytometry staining protocols.</li>
	<li>Centrifuge cells at 4 &#176;C, 300 x g for 5 min and resuspend cells in 30 &#181;L 1x PBS. Count cells and determine the viability via trypan blue staining.</li>
	<li>Determine the percentage of reverse transmigrated cells by using the following equation: 
	<img alt="Equation 4" src="/files/ftp_upload/56293/56293eq4.jpg" /> 
	NOTE: Accounting for the total number of forward transmigrated cells gives a more accurate indication of reverse transmigration as it is unlikely forward transmigration will be 100%.</li>
	<li>For analysis by microscopy, fix the gels by adding 100 &#181;L 2% formaldehyde (final concentration) to each well, cover the plates in aluminum foil and store at 4 &#176;C until analysis. For flow cytometry, do not fix the cells at this stage. See the protocols below for microscopy (section 6) or flow cytometry (section 7) analysis. 
	CAUTION: Formaldehyde is toxic and corrosive; use personal protective equipment (PPE). Care should be taken when adding formaldehyde, however, this step does not need to be performed in a fume hood due to the low concentration used.</li>
</ol>
6. Quantitation of Foam Cells and Macrophages by Microscopy (Oil-Red O Stain)
<ol>
	<li>Remove the formaldehyde and wash the gels with 100 &#181;L of 50% (v/v) methanol for 5 min and then 100 &#181;L 78% (v/v) methanol for 15 min. 
	NOTE: The staining steps may be performed on the laboratory bench and not in a Class II Biohazard cabinet.</li>
	<li>Stain for lipid droplets by adding 100 &#181;L of freshly prepared 0.2% (w/v) Oil-Red O (see Table of Materials for details) for 1 h at room temperature.</li>
	<li>Remove excess stain by washing the gels 4 times with 100 &#181;L of 78% (v/v) methanol, aspirating and discarding the supernatant after each wash.</li>
	<li>Counterstain for 15 min at room temperature with 100 &#181;L of Giemsa, diluted 1:10 in distilled water.</li>
	<li>Aspirate the Giemsa stain and wash gels once with water.</li>
	<li>To detach the gels from the plate, rim the wells using a 21 G needle, with the bevel facing outwards, around the edge of the gel.</li>
	<li>To mount the gels on a microscope slide, punch two holes (6.35 mm diameter) in a 2.54 cm x 1.5 cm strip of double-sided tape. Fix the tape to a standard microscope side and remove the protective coating from the top.
	<ol>
		<li>Add a drop of water to the holes in the tape using a transfer pipette. Using tweezers, gently transfer gels to the holes in the tape and cover with a size 1.5 glass coverslip. Press gently on the coverslip to adhere it to the tape.</li>
	</ol>
	</li>
	<li>Examine with differential interference contrast bright-field microscopy using 40X objective on an inverted microscope.
	<ol>
		<li>Bring the HUVEC monolayer into focus at 40X magnification and scroll &#39;into&#39; the gel, counting macrophages/foam cells throughout the entire depth of the gel. Repeat for three distinct fields of view located at similar distances from the edge of the gel in order to minimize possible edge effects. This will ensure gel depth is consistent between counting regions. 
		NOTE: Score cells in the gel as either foam cells (defined as cells containing &#62;1/3 of their cytoplasm as Oil-Red O stained lipid droplets) or macrophages within 2 h of mounting on slides (Figure 2). Foam cell formation is expressed as the percentage of foam cells relative to the total number of migrated cells and is expressed as the median counts in 3 fields of view examined for 6 gels per condition, therefore a median of 18 measurements per condition. An example of raw cell counts from a typical experiment is shown in Table 2. 
		NOTE: Classifying a cell as a foam cell vs macrophage by microscopy is subjective and consequently can be investigator-dependent. In clinical studies, where experiments are performed over an extended period of time, it is important for all counting to be performed blinded by a single investigator. Examples of foam cells and macrophages in gels are provided in Figure 2.</li>
	</ol>
	</li>
</ol>
7. Analysis of Transmigrated Cells by Flow Cytometry
<ol>
	<li>To phenotypically characterize foam cells and macrophages following transendothelial migration, digest the gels by adding 100 &#181;L of 37 &#176;C prewarmed 1 mg/mL collagenase D diluted in M199 to each well and incubate for 20 min at 37 &#176;C/5% CO2. 
	NOTE: Place the 96-well plates directly on clean metal racks in the incubator to ensure even heat distribution.</li>
	<li>Following incubation, macerate the gels using a 200 &#181;L pipette tip and incubate for a further 20 min at 37 &#176;C/5% CO2.</li>
	<li>Once fully digested, filter and pool the digested replicate gels through 35 &#181;m nylon mesh capped FACS tubes placed on ice, and wash once with FACS wash (see Table of Materials) at 4 &#176;C, 300 x g. Resuspend the cells in 100 &#181;L of FACS wash.</li>
	<li>Incubate the resulting cells with fluorophore-conjugated antibodies specific for CD45, a live/dead cell marker and surface/intracellular phenotypic or functional markers of interest using standard flow cytometry protocols. 
	NOTE: Prepare control tubes for compensation using the appropriate fluorophores and Ig compensation beads. Extracted cells may also be collected onto glass slides for immunofluorescence microscopy by cytospinning. Alternatively, we have successfully collected cells using this protocol for assessment via imaging flow cytometry.</li>
	<li>Acquire cells using a flow cytometer, and perform compensation as required. 
	NOTE: As foam cells/macrophages may not form individual populations using light scatter properties and may differ considerably in size and shape, set liberal forward scatter (FSC) and side scatter (SSC) gates in order to capture all migrated cells as shown in Figure 3A.</li>
	<li>Following acquisition, perform the data analysis using flow cytometry software. To identify migrated cells, first select cells as negative for the live/dead marker as shown in Figure 3B. Next, perform a single-cell discrimination by creating a tight gate around the cells shown on a SSC-area vs SSC-height plot.</li>
	<li>Select CD45+ cells and gate the major population of migrated cells present in a FSC/SSC plot as these are the macrophages/foam cells.</li>
</ol>

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The authors have nothing to disclose.

The protocol described here offers a versatile and physiologically relevant method for assessing the atherogenicity of monocytes from human clinical cohorts, by combining both monocyte transmigration and foam cell formation. This model offers advantages over alternative methods of foam cell formation as it takes into account the effect of monocyte transmigration on foam cell formation and allows the measurement of reverse transmigration6 in addition to the inherent propensity of monocytes to matur...

Monocyte transmigration is a crucial step in the development of atherosclerotic plaque that may lead to thrombosis, stroke and myocardial infarction. Atherosclerotic plaques develop from fatty streaks, generally present at sites of low oscillatory blood flow in medium to large arteries, where deposited lipid contributes to endothelial activation and localized inflammation1. Monocytes are recruited to endothelial cells in fatty streaks via monocyte chemotactic proteins (such as CCL2) and transmigrate into the intima2. Following transmigration, monocytes may form atherogenic, lipid-laden macrophages called foam cells as a consequence of lipid uptake, lipid synthesis, down-regulation of cholesterol efflux or a combination of the above factors. Monocytes may also accumulate lipids in the circulation and have a &#39;foamy&#39; phenotype, possibly predisposing cells for foam cell formation3,4. Foam cells are the defining feature of fatty streaks and early-stage atherosclerotic plaques and their formation is influenced by both lipid and inflammatory mediators5. Alternatively, monocytes have the ability to reverse transmigrate from the artery into the bloodstream6, thereby removing lipid from the intima and acting to maintain the health of the artery.
Determining the propensity of monocytes to transmigrate across arterial endothelium and form foam cells in the intima, or to reverse transmigrate and carry lipid out of the plaque, is a key requirement for understanding the role of monocyte activation in increasing atherosclerotic risk. Mouse models of CADs such as atherosclerosis are important in elucidating real-time in vivo information on fatty streak/atherosclerotic plaque development. However, these models require a genetic alteration of the natural cholesterol processing abilities of these animals usually coupled with drastic alterations in diet (such as the ApoE-/- Western-type diet model)7,8, thereby, inducing non-physiological accumulation of circulating lipid levels which drive plaque development. These models may have limited relevance to chronic inflammatory human conditions such as HIV infection which are not associated with increased circulating cholesterol or low-density lipoprotein (LDL) levels. Furthermore, differences in monocyte biology between humans and mice make the testing of immunological questions regarding the relevance of subpopulations of monocytes (such as intermediate monocytes (CD14++CD16+))9 difficult. This is important when studying the mechanisms driving cardiovascular disease as intermediate monocyte counts independently predict cardiovascular events10,11. While assays exist to sequentially measure either monocyte transmigration or foam cell formation in isolation, no in vitro assay has been validated for quantifying both aspects of early atherogenesis using the same cells from clinical cohorts. Transwell models utilize a modified Boyden two-chamber system whereby cells are loaded into the top chamber and transmigrate across a porous plastic barrier or cell monolayer into a lower chamber that typically contains media with chemoattractant12,13. Whilst widely used for analyzing leukocyte transmigration, these models do not generally incorporate a layer representing the intima, resulting in transmigrated cells migrating into solution, and do not allow for the measurement of foam cell formation or reverse transmigration of the same cells. Conversely, models of foam cell formation do not account for any transmigratory-induced changes to monocytes or effects of endothelial activation which is known to contribute to foam cell formation14. Furthermore, these systems induce foam cell formation from macrophages adhered to cell culture plates by the addition of saturating concentrations of exogenous oxidized low-density lipoprotein (oxLDL)15,16, a key inducer of foam cell formation. LDL used in these models is often oxidized by non-physiologically-relevant processes such as CuSO4 treatment17, therefore, questioning the physiological importance of studies using these models.
Here we describe an assay that quantifies monocyte transmigration and foam cell formation of the same cells which does not require the addition of exogenous oxLDL, thus better modelling the role of monocytes in foam cell formation. This model was originally developed by Professor William Muller (Northwestern University, Chicago)18, and has been further refined in our laboratory to assess ex vivo the atherogenicity of monocytes isolated under non-activating conditions from individuals with underlying inflammatory conditions accompanying diseases such as HIV infection19 as well as ageing20, that are associated with an increased risk of atherosclerosis. This model also provides a platform for answering basic biological questions regarding the propensity of different monocyte subsets to form foam cells20, the influence of endothelial activation by cytokines such as TNF on foam cell formation14, and the migratory properties of monocytes such as the depth and speed of transmigration in gels19. Furthermore, monocyte transmigration and foam cell formation can be quantified using standard microscopy, live cell imaging, flow cytometry and imaging flow cytometry, therefore, providing a versatile method to evaluate the role of monocytes in atherogenesis.

<table><tbody><tr><td>&lt;strong&gt;Gel preparation reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>NaOH</td><td>Sigma-Aldrich</td><td>221465-500G</td><td>0.1 M NaOH diluted in H&lt;sub&gt;2&lt;/sub&gt;0</td></tr><tr><td>10x M199</td><td>Sigma-Aldrich</td><td>M0650</td><td></td></tr><tr><td>AcCOOH</td><td>Sigma-Aldrich</td><td>695092-100ML</td><td>20 mM Acetic acid diluted in H&lt;sub&gt;2&lt;/sub&gt;0</td></tr><tr><td>Cultrex Bovine Collagen I</td><td>R&amp;amp;D Systems</td><td>3442-050-01</td><td>Type I Fibrous Collagen</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Cell culture&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>M199</td><td>Life Technologies</td><td>11150-059</td><td>M199 media containing Earle&#039;s salts, L-glutamine and 2.2 g/L Sodium Bicarbonate.&lt;br/&gt; Media supplemented with 100 &amp;micro;g/mL L-glutamine&amp;nbsp; and 100 U/mL penicillin/streptomycin&amp;nbsp;</td></tr><tr><td>M20</td><td></td><td></td><td>Supplemented M199 containing 20% heat-inactivated pooled or donor serum.&lt;br/&gt; Individual lipid species such as LDL can be added to M199.</td></tr><tr><td>HUVEC</td><td></td><td></td><td>Primary human umbilical cord endothelial cells (HUVEC) can be isolated from umbilical cords donated with informed consent and ethics approval. Isolated HUVEC may also be purchased commercially.</td></tr><tr><td>Human coronary artery endothelial cells</td><td></td><td></td><td>Primary human coronary artery endothelial cells can be isolated from arteries donated with informed consent and ethics approval. Isolated cells may also be purchased commercially.</td></tr><tr><td>EDTA</td><td>BDH Merck</td><td>10093.5V</td><td>Ethylenediaminetetraacetic acid (EDTA) - 0.5 M, pH 8.0</td></tr><tr><td>EGTA</td><td>Sigma-Aldrich</td><td>E3889-500G</td><td>Ethylene glycol-bis(&amp;beta;-aminoethyl ether)-N,N,N&#039;,N&#039;-tetraacetic acid (EGTA) - 1 mM, pH 8.0</td></tr><tr><td>0.05% trypsin EDTA</td><td>Gibco</td><td>25300-054</td><td>0.05% trypsin/0.53 EDTA (1X)</td></tr><tr><td>L-glutamine</td><td>Gibco</td><td>25030-081</td><td>L-glutamine (200 mM)</td></tr><tr><td>Penicillin/streptomycin&amp;nbsp;</td><td>Gibco</td><td>15140-122</td><td>Penicillin/streptomycin (10,000 units/mL Penicillin and 10,000 &amp;micro;g/mL Streptomycin)</td></tr><tr><td>New born calf serum</td><td>Gibco</td><td>16010-142</td><td>New born calf serum: New Zealand origin</td></tr><tr><td>Fibronectin</td><td>Sigma-Aldrich</td><td>F1056-1MG</td><td>Fibronectin - 50 &amp;micro;g/mL aliquots prepared and stored</td></tr><tr><td>PBS (1x)</td><td>Gibco</td><td>14200-075</td><td>Dulbecco&#039;s Phosphate Buffered Saline (10x): Dilute to 1x with sterile H&lt;sub&gt;2&lt;/sub&gt;0</td></tr><tr><td>0.1% TNF</td><td>Gibco</td><td>PHC3015</td><td>Recombinant human TNF - Reconstituted in H20 and stored in 10 &amp;micro;g/mL aliquots</td></tr><tr><td>Low-density lipoprotein</td><td>Merck Millipore</td><td>LP2-2MG</td><td>Low-density lipoprotein (LDL)</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Microscopy&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>1 or 2% formaldehyde</td><td>Polysciences</td><td>4018</td><td>1 or 2% formaldehyde diluted with sterile H&lt;sub&gt;2&lt;/sub&gt;0</td></tr><tr><td>50% and 78% methanol</td><td>Ajax Finechem</td><td>318-2.5L GL</td><td>50% or 78% v/v methanol, diluted with H&lt;sub&gt;2&lt;/sub&gt;0</td></tr><tr><td>Oil Red O stain</td><td>Sigma-Aldrich</td><td>O0625-25G</td><td>Dilute to 2 mg/mL in 22% 1M NaOH and 78% (v/v) methanol</td></tr><tr><td>Microscope slides</td><td>Mikro-Glass</td><td>S41104AMK</td><td>Twin frosted 45 degree ground edge microscope slides (25 X 76 mm)</td></tr><tr><td>Cover slips</td><td>Menzel-Gl&amp;auml;ser</td><td>MENCS224015GP</td><td>22 x 40 mm #1.5 size glass cover slips</td></tr><tr><td>Double-sided tape</td><td>3M Scotch</td><td>4011</td><td>Super strength exterior mounting tape (25.4 mm x 1.51 m)</td></tr><tr><td>Giemsa stain</td><td>Merck Millipore</td><td>1.09204.0500</td><td>Giemsa&#039;s azur eosin methylene blue solution (dilute stock 1:10 in H&lt;sub&gt;2&lt;/sub&gt;0)</td></tr><tr><td>Hole punch</td><td></td><td></td><td>Hand-held single hole punch (6.35 mm punch)</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Flow cytometry&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Collagenase D</td><td>Roche Diagnostics</td><td>11088858001</td><td>Collagenase D diluted in M199 media to 1 mg/mL&amp;nbsp;</td></tr><tr><td>35 &amp;micro;m nylon mesh capped polystyrene FACS tubes</td><td>BD Biosciences</td><td>352235</td><td>35 &amp;micro;m nylon mesh capped polystyrene FACS tubes</td></tr><tr><td>Live/Dead Fixable Yellow Dead Cell stain</td><td>Life Technologies</td><td>L34959</td><td>Live/Dead Fixable Yellow Dead Cell stain</td></tr><tr><td>FACS wash</td><td></td><td></td><td>Prepare by mixing 1 X PBS&lt;sup&gt;-&lt;/sup&gt;, 2 mM EDTA and 1% New born calf serum</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Plasticware&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>96 well plate</td><td>Nunclon</td><td>167008</td><td>Delta surface flat-bottomed 96 well plate</td></tr><tr><td>10 cm Petri Dish</td><td>TPP</td><td>93100</td><td>Sterile 10 cm Petri dish</td></tr><tr><td>1.5 mL Eppendorf tubes</td><td>Eppendorf</td><td>0030 125.150</td><td>Eppendorf tubes</td></tr><tr><td>Transfer pipette</td><td>Samco Scientific</td><td>222-20S</td><td>Sterile transfer pipette (1 mL, large bulb)</td></tr></tbody></table>

quantification of monocyte transmigration and foam cell formation from individuals with chronic inflammatory conditions

Coronary artery disease (CAD) is a leading cause of morbidity and mortality worldwide. Atherosclerosis, a leading cause of CAD, is initiated by the transmigration of innate immune monocytes to inflammatory sites of deposited lipid called fatty streaks, which are present in arterial walls of medium to large arteries. The key pathogenic feature of lesions at this early stage of atherosclerosis is the maturation of monocytes which migrate into arteries to form foam cells or lipid-laden macrophages. Considerable evidence supports the hypothesis that risk of atherosclerosis is increased by chronic inflammatory conditions accompanying diseases such as rheumatoid arthritis and HIV, as well as general ageing, and that this risk is predicted by monocyte activation. While mouse models provide a good platform to investigate the role of monocytes in atherogenesis in vivo, they require genetic alteration of natural cholesterol metabolism and drastic alteration of normal mouse diets, and have limited suitability for the study of atherogenic influences of human comorbid diseases. This motivated us to develop a human in vitro model to measure the atherogenic potential of monocytes isolated from individuals with defined disease states. Currently, human in vitro models are limiting in that they evaluate monocyte transmigration and foam cell formation in isolation. Here we describe a protocol in which monocytes isolated from patient blood transmigrate across human endothelial cells into a type 1 collagen matrix, and their propensity to mature into foam cells in the presence or absence of exogenous lipid is measured. The protocol has been validated for the use of human monocytes purified from individuals with HIV infection and elderly HIV uninfected individuals. This model is versatile and allows monocyte transmigration and foam cell formation to be evaluated using either microscopy or flow cytometry as well as allowing the assessment of atherogenic factors present in serum or plasma.

Quantifying monocyte transmigration
Monocytes are added to the model as described in Figure 1, and six gels are prepared for each condition. Monocytes for 6 gels per donor (i.e., 5.0 x 104 monocytes per gel 6 gels = 3.0 x 105 monocytes per donor) are resuspended to a final volume of 600 &#181;L of M199 media containing the required serum/isolated lipid...

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Quantification of Monocyte Transmigration and Foam Cell Formation from Individuals with Chronic Inflammatory Conditions

We describe a protocol to measure transmigration by monocytes across human endothelial monolayers and their subsequent maturation into foam cells. This provides a versatile method to assess the atherogenic properties of monocytes isolated from people with different disease conditions and to evaluate factors in blood which may enhance this propensity.

Coronary artery disease (CAD) is a leading cause of morbidity and mortality worldwide. Atherosclerosis, a leading cause of CAD, is initiated by the ...

quantification-monocyte-transmigration-foam-cell-formation-from

Research

JoVE Journal

Immunology and Infection

13.3K Views.  Burnet Institute. Beschrijven we een protocol voor het meten van transmigratie door monocyten over menselijke endothelial monolayers en hun daaropvolgende rijping in schuim cellen. Dit biedt een veelzijdige methode te beoordelen van de atherogene eigenschappen van monocyten geïsoleerd uit mensen met verschillende ziekten en te evalueren van factoren in het bloed die deze neiging kunnen versterken.

Video: Quantification of Monocyte Transmigration and Foam Cell Formation from Individuals with Chronic Inflammatory Conditions

On-Chip Endothelial Inflammatory Phenotyping

Atherogenesis is potentiated by metabolic abnormalities that contribute to a heightened state of systemic inflammation resulting in endothelial dysfunction. However, early functional changes in endothelium that signify an individual's level of risk are not directly assessed clinically to help guide therapeutic strategy. Moreover, the regulation of inflammation by local hemodynamics contributes to the non-random spatial distribution of atherosclerosis, but the mechanisms are difficult to delineate in vivo. We describe a lab-on-a-chip based approach to quantitatively assay metabolic perturbation of inflammatory events in human endothelial cells (EC) and monocytes under precise flow conditions. Standard methods of soft lithography are used to microfabricate vascular mimetic microfluidic chambers (VMMC), which are bound directly to cultured EC monolayers.1 These devices have the advantage of using small volumes of reagents while providing a platform for directly imaging the inflammatory events at the membrane of EC exposed to a well-defined shear field. We have successfully applied these devices to investigate cytokine-,2 lipid-3, 4 and RAGE-induced5 inflammation in human aortic EC (HAEC). Here we document the use of the VMMC to assay monocytic cell (THP-1) rolling and arrest on HAEC monolayers that are conditioned under differential shear characteristics and activated by the inflammatory cytokine TNF-&alpha;. Studies such as these are providing mechanistic insight into atherosusceptibility under metabolic risk factors.

Microfluidic flow chambers etched by photolithography and fabricated from PDMS are applied to probe functional outcomes associated with EC dysfunction and inflammation. In a representative experiment, the ability of differential shear stress to modulate monocytic cell adhesion to cytokine activated EC monolayers is demonstrated.

Atherogenesis is potentiated by metabolic abnormalities that contribute to a heightened state of systemic inflammation resulting in endothelial ...

Bioengineering

An In vitro Model to Study Heterogeneity of Human Macrophage Differentiation and Polarization

Monocyte-derived macrophages represent an important cell type of the innate immune system. Mouse models studying macrophage biology suffer from the phenotypic and functional differences between murine and human monocyte-derived macrophages. Therefore, we here describe an in vitro model to generate and study primary human macrophages. Briefly, after density gradient centrifugation of peripheral blood drawn from a forearm vein, monocytes are isolated from peripheral blood mononuclear cells using negative magnetic bead isolation. These monocytes are then cultured for six days under specific conditions to induce different types of macrophage differentiation or polarization. The model is easy to use and circumvents the problems caused by species-specific differences between mouse and man. Furthermore, it is closer to the in vivo conditions than the use of immortalized cell lines. In conclusion, the model described here is suitable to study macrophage biology, identify disease mechanisms and novel therapeutic targets. Even though not fully replacing experiments with animals or human tissues obtained post mortem, the model described here allows identification and validation of disease mechanisms and therapeutic targets that may be highly relevant to various human diseases.

An In vitro Model to Study Heterogeneity of Human Macrophage Differentiation and Polarization

Monocyte-derived macrophages are important cells of the innate immune system. Here, we describe an easy to use in vitro model to generate these cells. Using gradient centrifugation, negative bead isolation and specific cell culture conditions, monocyte-derived macrophages can be generated for phenotypic and functional studies.

Monocyte-derived macrophages represent an important cell type of the innate immune system. Mouse models studying macrophage biology suffer from the ...

Isolation of Human Monocytes by Double Gradient Centrifugation and Their Differentiation to Macrophages in Teflon-coated Cell Culture Bags

Human macrophages are involved in a plethora of pathologic processes ranging from infectious diseases to cancer. Thus they pose a valuable tool to understand the underlying mechanisms of these diseases. We therefore present a straightforward protocol for the isolation of human monocytes from buffy coats, followed by a differentiation procedure which results in high macrophage yields. The technique relies mostly on commonly available lab equipment and thus provides a cost and time effective way to obtain large quantities of human macrophages. Briefly, buffy coats from healthy blood donors are subjected to a double density gradient centrifugation to harvest monocytes from the peripheral blood. These monocytes are then cultured in fluorinated ethylene propylene (FEP) Teflon-coated cell culture bags in the presence of macrophage colony-stimulating factor (M-CSF). The differentiated macrophages can be easily harvested and used for subsequent studies and functional assays. Important methods for quality control and validation of the isolation and differentiation steps will be highlighted within the protocol. In summary, the protocol described here enables scientists to routinely and reproducibly isolate human macrophages without the need for cost intensive tools. Furthermore, disease models can be studied in a syngeneic human system circumventing the use of murine macrophages.

We present a simple and efficient protocol for the generation of human macrophages. Buffy coats are processed by double density gradient centrifugation and isolated monocytes are then differentiated to macrophages in Teflon-coated cell culture bags. This maximizes macrophage yields and facilitates cell harvesting for subsequent experiments.

Human macrophages are involved in a plethora of pathologic processes ranging from infectious diseases to cancer. Thus they pose a valuable tool to ...

Culture of Macrophage Colony-stimulating Factor Differentiated Human Monocyte-derived Macrophages

A protocol is presented for cell culture of macrophage colony-stimulating factor (M-CSF) differentiated human monocyte-derived macrophages. For initiation of experiments, fresh or frozen monocytes are cultured in flasks for 1 week with M-CSF to induce their differentiation into macrophages. Then, the macrophages can be harvested and seeded into culture wells at required cell densities for carrying out experiments. The use of defined numbers of macrophages rather than defined numbers of monocytes to initiate macrophage cultures for experiments yields macrophage cultures in which the desired cell density can be more consistently attained. Use of cryopreserved monocytes reduces dependency on donor availability and produces more homogeneous macrophage cultures.

A protocol is presented for cell culture of macrophage colony-stimulating factor (M-CSF) differentiated human monocyte-derived macrophages. The protocol utilizes cryopreservation of monocytes coupled with their bulk differentiation into macrophages. Then harvested macrophages can then be seeded into culture wells at required cell densities for carrying out experiments.

A protocol is presented for cell culture of macrophage colony-stimulating factor (M-CSF) differentiated human monocyte-derived macrophages. For initiation ...

Quantifying Human Monocyte Chemotaxis In Vitro and Murine Lymphocyte Trafficking In Vivo

Chemotaxis is migration along a specific chemical gradient1. Chemokines are chemotactic cytokines that promote cellular trafficking with anatomic and temporal specificity2. Chemotaxis is a critical function of lymphocytes and other immune cells that can be quantitatively assessed in vitro. This manuscript describes methods that permit the evaluation of chemotaxis, both in vitro and in vivo, for diverse cell types including cell lines and native cells. The in vitro, plate-based format permits the comparison of several conditions simultaneously in real-time, and can be completed within 1-4 h. In vitro assay conditions can be manipulated to introduce agonists and antagonists, as well as differentiate chemotaxis from chemokinesis, which is random movement. For in vivo trafficking assessments, immune cells can be labeled with multiple fluorescent dyes and used for adoptive transfer. The differential labeling of cells allows for mixed cell populations to be introduced into the same animal, thereby decreasing variance and reducing the number of animals required for an adequately powered experiment. Migration into lymphoid tissue occurs in as little as 1 h, and multiple tissue compartments can be sampled. Flow cytometry following tissue harvest allows for a rapid and quantitative analysis of the migratory patterns of multiple cell types.

Quantifying Human Monocyte Chemotaxis In Vitro and Murine Lymphocyte Trafficking In Vivo

Protocols for quantitative assessment of lymphocyte chemotaxis and migration are important tools for immunology research. Here, an in vitro protocol is described that permits real-time, multiplexed evaluation of cell migration, as well as a complementary in vivo technique enabling tracking of native cells to spleen.

Chemotaxis is migration along a specific chemical gradient1. Chemokines are chemotactic cytokines that promote cellular trafficking with anatomic and ...

Laminar Flow-based Assays to Investigate Leukocyte Recruitment on Cultured Vascular Cells and Adherent Platelets

The recruitment of leukocytes upon injury or inflammation to sites of injury or tissue damage has been investigated during recent decades and has resulted in the concept of the leukocyte adhesion cascade. However, the exact molecular mechanisms involved in leukocyte recruitment have not yet been fully identified. Since leukocyte recruitment remains an important subject in the field of infection, inflammation, and (auto-) immune research, we present a straightforward laminar flow-based assay to study underlying mechanisms of the adhesion, de-adhesion, and transmigration of leukocytes under venous and arterial flow regimes. The in vitro assay can be used to study the molecular mechanisms that underlie the interactions between leukocytes and their cellular partners in different models of vascular inflammation. This protocol describes a laminar flow-based assay using a parallel-flow chamber and an inverted phase contrast microscope connected to a camera to study the interactions of leukocytes and endothelial cells or platelets, which can be visualized and recorded then analyzed offline. Endothelial cells, platelets, or leukocytes can be pretreated with inhibitors or antibodies to determine the role of specific molecules during this process. Shear conditions, i.e. arterial or venous shear stress, can be easily adapted by the viscosity and flow rate of the perfused fluids and the height of the channel.

Leukocytes avidly interact with vascular cells and platelets after vessel wall injury or during inflammation. Here, we describe a straightforward laminar flow-based assay to characterize the molecular mechanisms that underlie the interactions between leukocytes and their cellular partners.

The recruitment of leukocytes upon injury or inflammation to sites of injury or tissue damage has been investigated during recent decades and has resulted ...

Simultaneous Study of the Recruitment of Monocyte Subpopulations Under Flow In Vitro

The recruitment of monocytes from the blood to targeted peripheral tissues is critical to the inflammatory process during tissue injury, tumor development and autoimmune diseases. This is facilitated through a process of capture from free flow onto the luminal surface of activated endothelial cells, followed by their adhesion and transendothelial migration (transmigration) into the underlying affected tissue. However, the mechanisms that support the preferential and context-dependent recruitment of monocyte subpopulations are still not fully understood. Therefore, we have developed a method that allows the recruitment of different monocyte subpopulations to be simultaneously visualized and measured under flow. This method, based on time-lapse confocal imaging, allows for the unambiguous distinction between adherent and transmigrated monocytes. Here, we describe how this method can be used to simultaneously study the recruitment cascade of pro-angiogenic and non-angiogenic monocytes in vitro. Furthermore, this method can be extended to study the different steps of recruitment of up to three monocyte populations.

Here, we present an integrated protocol that measures monocyte subpopulation trafficking under flow in vitro by use of specific surface markers and confocal fluorescence microscopy. This protocol can be used to explore sequential recruitment steps as well as to profile other leukocyte subtypes using other specific surface markers.

The recruitment of monocytes from the blood to targeted peripheral tissues is critical to the inflammatory process during tissue injury, tumor development ...

Fluorescence-mediated Tomography for the Detection and Quantification of Macrophage-related Murine Intestinal Inflammation

Murine models of disease are indispensable to scientific research. However, many diagnostic tools such as endoscopy or tomographic imaging are not routinely employed in animal models. Conventional experimental readouts often rely on post mortem and ex vivo analyses, which prevent intra-individual follow-up examinations and increase the number of study animals needed. Fluorescence-mediated tomography enables the non-invasive, repetitive, quantitative, three-dimensional assessment of fluorescent probes. It is highly sensitive and permits the use of molecular makers, which allows for the specific detection and characterization of distinct molecular targets. In particular, targeted probes represent an innovative tool for analyzing gene activation and protein expression in inflammation, autoimmune disease, infection, vascular disease, cell migration, tumorigenesis, etc. In this article, we provide step-by-step instructions on this sophisticated imaging technology for the in vivo detection and characterization of inflammation (i.e., F4/80-positive macrophage infiltration) in a widely used murine model of intestinal inflammation. This technique might also be used in other research areas, such as immune cell or stem cell tracking.

Target-specific probes represent an innovative tool for analyzing molecular mechanisms, such as protein expression in various types of disease (e.g., inflammation, infection, and tumorigenesis). In this study, we describe a quantitative three-dimensional tomographic assessment of intestinal macrophage infiltration in a murine model of colitis using F4/80-specific fluorescence-mediated tomography.

Murine models of disease are indispensable to scientific research. However, many diagnostic tools such as endoscopy or tomographic imaging are not ...

Medicine

Quantification of Monocyte Chemotactic Activity In Vivo and Characterization of Blood Monocyte Derived Macrophages

Tissue homeostasis and repair are critically dependent on the recruitment of monocyte-derived macrophages. Both under- and over-recruitment of monocyte-derived macrophages can impair wound healing. We showed that high fat and high sugar diets promote monocyte priming and dysfunction, converting healthy blood monocytes into a hyper chemotactic phenotype poised to differentiate into macrophages with dysregulated activation profiles and impaired phenotypic plasticity. The over-recruitment of monocyte-derived macrophages and recruitment of macrophages with dysregulated activation profiles is believed to be a major contributor to the development of chronic inflammatory diseases associated with metabolic disorders, including atherosclerosis and obesity. The goal of this protocol is to quantify the chemotactic activity of blood monocytes as a biomarker for monocyte priming and dysfunction and to characterize the macrophage phenotype blood monocytes are poised to differentiate into in these mouse models. Using single cell Western blot analysis, we show that after 24 h 33%of cells recruited into MCP-1-loaded basement membrane-derived gel plugs injected into mice are monocytes and macrophages; 58% after day 3. However, on day 5, monocyte and macrophage numbers were significantly decreased. Finally, we show that this assays also allows for the isolation of live macrophages from the surgically retrieved basement membrane-derived gel plugs, which can then be subjected to subsequent characterization by single cell Western blot analysis.

Here we present a protocol to quantify the chemotactic activity of blood monocytes in mouse models, to assess the effects of nutritional, pharmacological and genetic interventions on blood monocyte and to characterize the blood monocytes derived macrophages in mouse models using monocyte-chemoattractant protein-1 (MCP-1)-loaded basement membrane-derived gel plugs.

Tissue homeostasis and repair are critically dependent on the recruitment of monocyte-derived macrophages. Both under- and over-recruitment of ...

Flow Cytometry Analysis of Immune Cell Subsets within the Murine Spleen, Bone Marrow, Lymph Nodes and Synovial Tissue in an Osteoarthritis Model

Osteoarthritis (OA) is one of the most prevalent musculoskeletal diseases, affecting patients suffering from pain and physical limitations. Recent evidence indicates a potential inflammatory component of the disease, with both T-cells and monocytes/macrophages potentially associated with the pathogenesis of OA. Further studies postulated an important role for subsets of both inflammatory cell lineages, such as Th1, Th2, Th17, and T-regulatory lymphocytes, and M1, M2, and synovium-tissue-resident macrophages. However, the interaction between the local synovial and systemic inflammatory cellular response and the structural changes in the joint is unknown. To fully understand how T-cells and monocytes/macrophages contribute towards OA, it is important to be able to quantitively identify these cells and their subsets simultaneously in synovial tissue, secondary lymphatic organs and systemically (the spleen and bone marrow). Nowadays, the different inflammatory cell subsets can be identified by a combination of cell-surface markers making multi-color flow cytometry a powerful technique in investigating these cellular processes. In this protocol, we describe detailed steps regarding the harvest of synovial tissue and secondary lymphatic organs as well as generation of single cell suspensions. Furthermore, we present both an extracellular staining assay to identify monocytes/macrophages and their subsets as well as an extra- and intra-cellular staining assay to identify T-cells and their subsets within the murine spleen, bone marrow, lymph nodes and synovial tissue. Each step of this protocol was optimized and tested, resulting in a highly reproducible assay that can be utilized for other surgical and non-surgical OA mouse models.

Here, we describe a detailed and reproducible flow cytometry protocol to identify monocyte/macrophage and T-cell subsets using both extra- and intracellular staining assays within the murine spleen, bone marrow, lymph nodes and synovial tissue, utilizing an established surgical model of murine osteoarthritis.

Quantification of Monocyte Transmigration and Foam Cell Formation from Individuals with Chronic Inflammatory Conditions

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