This study was supported by grants from the intramural innovative medical research program of Muenster University medical faculty (KE121201 to C.K.) and the German Research Foundation (DFG, Fo354/3-1 to D.F.).

NOTE: Please refer to Supplemental Table 1 for preparation of buffers and stock solutions.
1. Protein expression in E. coli
<ol>
	<li>Cloning
	<ol>
		<li>Clone tag-free human S100A12 (NCBI Reference Sequence: NP_005612.1) into bacterial expression vector pET11b. To express the protein, transform the construct into E. coli BL21(DE3).</li>
	</ol>
	</li>
	<li>Culture
	<ol>
		<li>Prepare a starter culture by inoculating a single colony in 5 mL of growth medium (LB broth with 100 &#181;g/mL ampicillin) in a 14 mL round-bottom tube. Incubate overnight at 37 &#176;C with shaking at 220 rpm. Transfer 2&#8722;4 mL of overnight culture into 400 mL of growth medium in a 2 L Erlenmayer flask and incubate the culture at 37 &#176;C with shaking at 220 rpm. 
		NOTE: Initial density of the main culture should be optical density at 600 nm (OD600) = 0.1.</li>
		<li>Monitor the OD600 during growth. Induce protein expression by addition of 1 M isopropyl-&#223;-D-thiogalactopyranosid (IPTG) to a final concentration of 1 mM at OD600 = 0.5&#8722;0.6. Incubate at 37 &#176;C and 220 rpm for additional 4 h. 
		NOTE: In general, an OD600 of 0.6 will be reached after 1.5&#8722;2.5 h at 37 &#176;C.</li>
		<li>Prepare 50 mL sonication buffer by dissolving 50 mM Tris, 50 mM NaCl and 1 mM ethylenediamine tetraacetic acid (EDTA) in 40 mL of deionized water. Adjust pH with HCl to 8.0 and make up to 50 mL. Add protease inhibitor (1 tablet per 50 mL solution) and equilibrate the buffer to 4 &#176;C.</li>
		<li>Transfer the bacterial culture into suitable centrifuge bottles and harvest the cells at 3,200 x g for 30 min at 4 &#176;C. Discard the supernatant and resuspend the pellet in 25 mL of ice-cold sonication buffer. Henceforth keep the cells on ice. 
		NOTE: Resuspended cells can be stored at -20 &#176;C for short-term and at -80 &#176;C for long-term.</li>
	</ol>
	</li>
	<li>Sonication/lysis
	<ol>
		<li>Sonicate the cells for 6 cycles of 30 s on ice. After each cycle, rest cells for 30&#8722;60 s to protect the cells from overheating.</li>
		<li>Transfer the cell suspension to a pre-chilled 50 mL high-speed centrifugation tube and centrifuge in a fixed angle rotor at 15,000 x g for 30 min at 4 &#176;C. Decant the cleared lysate which contains the soluble cytosolic proteins into a fresh 50 mL tube and discard the pellet.</li>
	</ol>
	</li>
</ol>
2. Protein Purification
<ol>
	<li>Anion-exchange chromatography
	<ol>
		<li>Dialysis
		<ol>
			<li>Prepare anion-exchange chromatography (AIEX) buffer A by dissolving 20 mM Tris, 1 mM EDTA and 1 mM ethylene glycol-bis(2-aminoethylether)-N,N,N&#8217;,N&#8217;-tetraacetic acid (EGTA) in deionized water and adjust the pH to 8.5 with HCl. For dialysis prepare 2 x 5 L and for chromatography 2x 1 L of AIEX buffer A. 
			NOTE: The dialysate volume should be at about 100 times the sample volume. All buffers used for chromatography should be filtered (0.45 &#181;m or smaller) and degassed (e.g., by ultrasonic bath or vacuum degassing).</li>
			<li>Cut dialysis tubing (molecular weight cut-off [MWCO]: 3.5 kDa) into an appropriate length with additional space for air to ensure sample buoyancy above the rotating stir bar. 
			NOTE: Glycerol preserves the membrane and must be removed before use.</li>
			<li>To reduce the viscosity of the cleared protein solution from step 1.3.2, dilute the solution with 25 mL of AIEX buffer A to facilitate subsequent application to the chromatography column. Attach the first closure onto the tubing, load the sample into the membrane and attach the second closure at least 1&#160;cm from the top end of the tubing.</li>
			<li>Place the 5 L container with AIEX buffer A on a stir plate, add a stir bar and the membrane filled with protein solution. Adjust the speed to rotate the sample by avoiding interference with the rotating stir-bar. Dialyze for 12&#8722;24 h at 4 &#176;C, then replace the dialysate buffer (AIEX buffer A) by a fresh pre-cooled preparation and continue for at least 4 additional hours. Transfer the dialyzed protein solution to a 50 mL tube and filter through a 0.45 &#181;m filter unit. 
			NOTE: Storage possible.</li>
		</ol>
		</li>
		<li>Chromatography</li>
		<li>Start the liquid chromatography system (FPLC) with general maintenance, connect column buffers AIEX A and AIEX B (AIEX buffer A with 1 M NaCl) and the anion-exchange resin containing column. Refer to Table 1 for general chromatographic parameters. 
		NOTE: Buffers, column and FPLC equipment should be equilibrated to the same temperature before starting the run (refer to chromatographic parameters in Table 1, Table 2, Table 3, Table 4, and Table 5).</li>
		<li>Equilibrate the column with AIEX buffer A, subsequently load the sample onto the column and elute the proteins with a linear gradient from 0% to 100% high-salt buffer (AIEX B). Refer to Table 2 for a detailed method protocol.</li>
		<li>Collect 2 mL fractions during elution and analyze 10 &#181;L of each fraction on a Coomassie-stained 15% sodium dodecyl sulfate polyacrylamide&#160;gel&#160;electrophoresis (SDS-PAGE). Pool the fractions containing S100A12 protein for dialysis. 
		NOTE: The molecular weight of S100A12 is 10,575 Da.</li>
	</ol>
	</li>
	<li>Calcium-dependent hydrophobic-interaction chromatography (HIC)
	<ol>
		<li>Dialysis
		<ol>
			<li>Dialyze the protein solution against 20 mM Tris, 140 mM NaCl, pH 7.5 following the procedure described in section 2.1.1.</li>
		</ol>
		</li>
		<li>Chromatography
		<ol>
			<li>Prepare 1 L of chromatography buffer HIC A by dissolving 20 mM Tris, 140 mM NaCl and 25 mM CaCl2 in deionized water and adjust the pH to 7.5. For HIC buffer B, dissolve 20 mM Tris, 140 mM NaCl and 50 mM EDTA. Adjust the pH to 7.0 and filter and degas the buffers. Add CaCl2 to the sample to a final concentration of 25 mM and filter through 0.45 &#181;m. Equilibrate HIC buffers and sample to 4 &#176;C (column temperature).</li>
			<li>Start the liquid chromatography system with general maintenance, connect column buffers HIC A and B and the column. Refer to Table 3 for further chromatographic parameters.</li>
			<li>Equilibrate the column, load the sample and extend the &#8216;wash unbound sample&#8217; block until the UV signal reaches baseline level again. Then start elution with a calcium chelator containing buffer (EDTA). Refer to Table 4 for a detailed method protocol. 
			NOTE: Previous experiments have shown that an excess of calcium seems to be beneficial for binding of S100A12 to the chromatography resin.</li>
			<li>Collect peak fractions of 2 mL and analyze 10 &#181;L of each fraction on a Coomassie-stained 15% SDS-PAGE. Pool pure S100A12 fractions and dialyze against Hepes-buffered saline (HBS; 20 mM Hepes, 140 mM NaCl, pH 7.0) as described in section 2.1.1. 
			NOTE: Extinction coefficient of monomeric S100A12 is 2980 M-1 cm-1.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
3. Detection and Removal of Endotoxin
<ol>
	<li>Detection of endotoxin
	<ol>
		<li>To determine the endotoxin contamination, measure concentrations of diluted protein from step 2.2.2.4. (e.g., 1:10 and 1:100 in HBS) using an enzyme-linked immunosorbent assay (ELISA)-based, fluorescent endotoxin detection assay (Table of Materials). Perform this assay by following the manufacturer&#8217;s protocol. 
		NOTE: Use freshly prepared HBS solutions dissolved in ultrapure deionized water to avoid (new) endotoxin contamination by the buffer.</li>
	</ol>
	</li>
	<li>Removal of endotoxin and concentration of protein
	<ol>
		<li>Load 15 mL of sample onto a 50 kDa centrifugal filter unit and centrifuge at 3,200 x g and 10 &#176;C for approximately 10 min. Transfer the flow-through into a fresh vessel (on ice) and refill and centrifuge the 50 kDa filter tube as often as necessary. Wash the filter membrane twice with HBS to recover as much protein as possible after each step.</li>
		<li>Concentrate the S100A12-containing flow-through by using a 3 kDa centrifugal filter until the volume is reduced to one fifth up to one tenth of the initial loading volume (centrifugation at 3,200 x g, 10 &#176;C for approximately 30 min). Refill the filter as often as necessary, rinse the membrane and transfer the concentrated solution to a new tube after each refill. Discard the flow-through. Filter again through 50 kDa as described above. 
		NOTE: During this procedure, the loss of protein is remarkable (up to 50%), but the remaining protein preparation is completely depleted from LPS. This method yields about 10&#8722;15 mg protein from 400 mL culture.</li>
		<li>Adjust the protein solution to 1 mg/mL with endotoxin-free HBS and measure the LPS content as described in step 3.1.1. In case the protein solution is still not tested as LPS-free (&#60;0.1 EU/mL), eliminate remnant contaminations by using an endotoxin removal resin. 
		NOTE: With a protein concentration of 1 mg/mL, contamination of 0.1 EU/mL LPS equals approximately 0.01 pg LPS/&#181;g protein.</li>
	</ol>
	</li>
</ol>
4. Chemical Crosslinking and Oligomer Separation
<ol>
	<li>Chemical crosslinking
	<ol>
		<li>Prepare highly pure (endotoxin-free) stock solutions of 1 M CaCl2 and 100 mM ZnCl2 in ultrapure deionized water (Table of Materials). Use this buffer, freshly made, for the next step.</li>
		<li>Incubate 10 mL of purified endotoxin-free S100A12 (concentration 1 mg/mL in HBS) for 30 min at room temperature (RT) with either 25 mM CaCl2 for dimeric/tetrameric, or 25 mM CaCl2 and 1 mM ZnCl2 for hexameric/tetrameric S100A12 oligomers.</li>
		<li>Prepare crosslinker by dissolving 8 mg of BS3 in 500 &#181;L of endotoxin-free water directly before use (8 mg crosslinker for 10 mL ion-spiked protein solution equals a final concentration of 1.4 mM). Mix crosslinker and sample by pipetting and incubate for additional 30 min at RT. Quench the reaction by adding 1 M Tris-HCl, pH 7.5 to a final concentration of 50 mM and filter through 0.45 &#181;m.</li>
	</ol>
	</li>
	<li>Size-exclusion chromatography
	<ol>
		<li>Equilibrate the crosslinked sample to 12&#8722;15 &#176;C (column temperature) and start the liquid chromatography system with general maintenance. Connect column buffer (HBS) and the size-exclusion column. Refer to Table 5 for detailed information.</li>
		<li>Equilibrate the column in HBS, load sample and collect peak fractions (1&#8722;2 mL) during the run. Analyze these fractions on a 4&#8722;20% gradient SDS-PAGE and pool fractions with major bands of the desired protein complex. 
		NOTE: Hydrolysis of NHS ester reagents like BS3 in aqueous solutions results in a strong absorbance at 280 nm. Unbound crosslinker (molecular weight: 572 g/mol) elutes at the end of the run and results in a strong peak.</li>
		<li>Concentrate the solutions by using centrifugal filter units with MWCOs of 10 kDa (dimer), 30 kDa (tetramer) or 50 kDa (hexamer). Determine the endotoxin contamination as described in section 3.1. If necessary, remove remaining LPS with an endotoxin removal resin following the manufacturer&#8217;s recommendations (Table of Materials).</li>
	</ol>
	</li>
</ol>
5. Functional Testing on Monocytes
<ol>
	<li>Preparation of monocytes
	<ol>
		<li>Isolate monocytes from human buffy coats by density gradient centrifugation and subsequent monocyte enrichment by using a magnetic bead separation kit (Table of Materials). 
		NOTE: This protocol will result in approximately 5&#8722;7 x 107 monocytes (one buffy coat) with a purity of 83&#8722;95%. Since the number, but also the responsiveness of cells depends strongly on the donor, the protocol may have to be scaled up (depending on the required cell count).</li>
		<li>For density centrifugation, equilibrate the separation solution (density = 1.077 g/mL) to RT and transfer 20 mL into 50 mL centrifuge tubes (2 tubes per buffy coat). Dilute blood from the human buffy coat with Hank&#8217;s buffered salt solution (HBSS) to a total volume of 60 mL and layer 30 mL of this mixture carefully on top of the separation medium. Centrifuge at 550 x g for 35 min at RT. Disable the centrifuge brake.</li>
		<li>After centrifugation, the mononuclear peripheral blood cells (PBMCs) are located directly on top of the separation medium. Transfer these cells into a fresh 50 mL centrifuge tube, make up to 50 mL with HBSS, and centrifuge at 170 x g for 10 min. Aspirate the supernatant and resuspend the cell pellet in a small volume of HBSS by pipetting.</li>
		<li>Fill the tube up to 50 mL and centrifuge at 290 x g for 10 min. Aspirate the supernatant again, resuspend the cells in HBSS (50 mL) and centrifuge at 170 x g for 10 min. Count the cells and resuspend them in cell separation buffer (Table of Materials) to a concentration of 5 x 107 cells/mL. 
		NOTE: Instead of HBSS, phosphate-buffered saline (PBS) can be used for washing the cells.</li>
		<li>For monocyte isolation from PBMCs, use a magnetic negative cell isolation kit and follow the manufacturer&#8217;s protocol. Count monocytes and resuspend in monocyte medium (RPMI 1640, 15% heat-inactivated fetal calf serum [FCS], 4 mM L-glutamine, 100 U/mL penicillin/streptomycin) to a concentration of 2 x 106 cells/mL.</li>
		<li>To culture monocytes, coat culture dishes (e.g., 100 mm) with a hydrophobic, gas-permeable film, suitable for suspension cells (Table of Materials). Sterilize the plates by using UV light for approximately 30 min. Transfer the cells to these culture plates and let them rest over-night at 37 &#176;C and 5% CO2. 
		NOTE: Use 15&#8722;25 mL of cell suspension per coated dish.</li>
	</ol>
	</li>
	<li>Monocyte stimulation
	<ol>
		<li>Stimulation with S100A12 (wildtype) 
		NOTE: To distinguish untreated S100A12 (end-product from section 2.2.2) from crosslinked protein, S100A12 in the following is referred to as &#8216;wildtype&#8217;.
		<ol>
			<li>Transfer the rested cells into a 50 mL centrifugal tube and centrifuge at 350 x g for 10 min. Aspirate the supernatant and resuspend the cell pellet in stimulation medium (RPMI 1640, 5% heat-inactivated FCS, 4 mM L-glutamine, 100 U/mL penicillin/streptomycin) at a concentration of 2 x 106 cells/mL.</li>
			<li>For stimulation, use 24 well suspension plates and add 250 &#181;L of cell suspension per well (0.5 x 106 cells/well). Add 50 &#181;g/mL polymyxin B to the intended wells, followed by either LPS in different concentrations (25, 50, 100 and 200 pg/mL) or wildtype S100A12 (10, 20, 40, 60 &#181;g/mL). Further, apply the protein either untreated or heat-denatured (99 &#176;C, 10 min) in different concentrations to the cells. 
			NOTE: A short heat treatment denatures S100A12 protein but has less to no effect on LPS.</li>
			<li>Incubate plates for 4 h at 37 &#176;C and 5% CO2. Harvest the cells by transferring the cell suspension of each well to 1.5 mL reaction tubes. Centrifuge at 500 x g for 10 min. Transfer the supernatants to fresh tubes and measure TNF&#945; release in different dilutions (e.g., 1:2, 1:5, 1:10) with a human TNF&#945; ELISA kit following the manufacturer&#8217;s recommendations.</li>
		</ol>
		</li>
		<li>Stimulation with S100A12 oligomers
		<ol>
			<li>Prepare and seed out monocytes in 24 well suspension plates as described above. Stimulate cells by adding S100A12 oligomers from step 4.2.3. in different molar concentrations (125 nM, 250 nM, 500 nM, 1000 nM). 
			NOTE: In order to compare the abilities of the different oligomers to stimulate monocytes, oligomers were applied to the cells in comparable molar concentrations.</li>
			<li>Incubate for 4 h at 37 &#176;C and 5% CO2, harvest the cells and measure TNF&#945; release in the supernatants as described above.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Purification+and+partial+characterization+of+canine+S100A12.">Purification and partial characterization of canine S100A12.</a> Biochimie. 92 (12), 1914-1922 (2010).</li><li>Hung, K. W., Hsu, C. C., Yu, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Solution+structure+of+human+Ca2+-bound+S100A12.">Solution structure of human Ca2+-bound S100A12.</a> Journal of Biomolecular NMR. 57 (3), 313-318 (2013).</li><li>Endotoxin Removal. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Application Note - Sartorius Stedim Biotech. , (2010).</li><li>Hogasen, A. K. M., Abrahamsen, T. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Polymyxin-B+Stimulates+Production+of+Complement+Components+and+Cytokines+in+Human+Monocytes.">Polymyxin-B Stimulates Production of Complement Components and Cytokines in Human Monocytes.</a> Antimicrobial Agents and Chemotherapy. 39 (2), 529-532 (1995).</li><li>Valentinis, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+effects+of+polymyxin+B+on+human+dendritic+cells+maturation+-+The+role+of+I+kappa+B-alpha/NF-kappa+B+and+ERK1/2+pathways+and+adhesion.">Direct effects of polymyxin B on human dendritic cells maturation - The role of I kappa B-alpha/NF-kappa B and ERK1/2 pathways and adhesion.</a> Journal of Biological Chemistry. 280 (14), 14264-14271 (2005).</li><li>Teuber, M., Miller, I. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Selective+Binding+of+Polymyxin-B+to+Negatively+Charged+Lipid+Monolayers.">Selective Binding of Polymyxin-B to Negatively Charged Lipid Monolayers.</a> Biochimica Et Biophysica Acta. 467 (3), 280-289 (1977).</li></ol>

The authors have nothing to disclose.

In this protocol, we describe tag-free bacterial expression of human S100A12 and its purification as well as separation into different ion-induced oligomers for immune cell stimulation. Compared to published literature on S100A12 protein purification8,23,24, the use of high CaCl2 (25 mM) in hydrophobic-interaction chromatography is to our knowledge unique. Several protocols applying concentrations from 1 to 5 mM do pr...

S100A12 is a calcium binding protein which is predominantly produced by human granulocytes. The protein is overexpressed during (systemic) inflammation and its serum levels, particularly in (auto)inflammatory diseases such as systemic juvenile idiopathic arthritis (sJIA), familial Mediterranean fever (FMF) or Kawasaki disease (KD) can inform about disease activity and response to therapy. Depending on pattern recognition receptors (PRRs) such as toll-like receptors (TLRs), the innate immune system can be activated by pathogen-associated molecular patterns (PAMPs) like lipopolysaccharides (LPS) or damage associated molecular patterns (DAMPs; also termed &#8216;alarmins&#8217;). DAMPs are endogenous molecules such as cellular proteins, lipids or nucleic acids1. DAMP-functions are well described for the members of the calgranulin protein family, S100A8/A9 and S100A122, which are also reported to operate as divalent metal ion-chelating antimicrobial peptides3,4,5,6. Depending on the available ion strength S100A12 can, like other members of the S100 family, arrange into different homomultimers and until recently the impact of S100A12-oligomerisation on PRR-interaction, particularly TLR4, was unknown.
The protein&#8217;s monomeric form (92 amino acids, 10.2 kDa) consists of two EF-hand helix-loop-helix structures connected by a flexible linker. The C-terminal EF-hand contains the classical Ca2+-binding motif whereas the N-terminal EF-hand exhibits an S100 protein-specific extended loop structure (&#8216;pseudo-EF-hand&#8217;) and reveals reduced Ca2+-affinity. Ca2+-binding by S100A12 can induce a major conformational change in the proteins&#8217; C-terminus, which results in exposure of a hydrophobic patch on each monomer and forms the dimerization interface. Thus, under physiological conditions, the smallest quaternary structure formed by S100A12 is a non-covalent dimer (approximately 21 kDa) in which individual monomers are in antiparallel orientation. When arranged as dimer, S100A12 is reported to sequester Zn2+ as well as other divalent metal ions, e.g., Cu2+ with high affinity7. These ions are coordinated at the S100A12 dimer interface by amino acids H15 and D25 of one subunit and H85 as well as H89 of the anti-paralleling other subunit8,9,10. While earlier studies propose that Zn2+-loaded S100A12 may induce the protein&#8217;s organization into homo-tetramers (44 kDa) and to result in increased Ca2+-affinity11,12, recent metal titration studies6 suggest Ca2+-binding by S100A12 to increase the protein&#8217;s affinity to Zn2+. Once the S100A12 EF-hands are fully occupied by Ca2+, additional Ca2+ is thought to bind between dimers, triggering hexamer formation (approximately 63 kDa). The architecture of the hexameric quarternary structure is clearly different from that of the tetramer. It is proposed that the tetramer interface is disrupted to give rise to new dimer-dimer interfaces which benefits hexamer formation10. S100A12 is almost exclusively expressed by human granulocytes where it constitutes about 5% of all cytosolic protein13. In its DAMP function S100A12 was historically described as agonist of the multi-ligand receptor for advanced glycation end-products (RAGE), then termed extracellular newly identified RAGE-binding protein (EN-RAGE)14. Albeit we earlier reported biochemical S100A12-binding to both RAGE and TLR415, we recently demonstrated human monocytes to respond to S100A12 stimulation in a TLR4-dependent manner16. This requires arrangement of S100A12 into its Ca2+/Zn2+-induced hexameric quarternary structure16.
Here we describe a purification procedure for recombinant human S100A12 and its ion-induced oligomers for immune cell stimulations16,17. This is based on a two-step chromatography strategy, which initially includes an anion-exchange column to isolate and concentrate the protein and remove bulk contaminations (e.g., endotoxins/lipopolysaccharides)18. Ion-exchange chromatography resins separate proteins on the basis of different net surface charges. For acidic proteins like S100A12 (isoelectric point of 5.81), a buffer system with a pH of 8.5 and a strong anion-exchange resin leads to a good separation. Bound proteins were eluted with a high-salt buffer gradient. With an increase of ionic strength negative ions in the elution buffer compete with proteins for charges on the surface of the resin. Proteins individually elute depending on their net charge and in result of that, the buffers described herein allow to isolate and concentrate the overexpressed S100A12 protein. Due to negatively charged groups in lipopolysaccharides, these molecules also bind to anion-exchange resins. However, their higher net charge results in later elution in the applied high-salt gradient. The second step of the purification procedure has been introduced for polishing purposes. This makes use of the calcium binding ability of S100A12 and removes remaining impurities on a hydrophobic-interaction column. Calcium binding of S100A12 leads to a conformational change and an exposure of hydrophobic patches on the surface of the protein. On that condition, S100A12 interacts with the hydrophobic surface of the resin. Upon calcium-chelating by EDTA, this interaction is reversed. In the presence of ions, especially calcium and zinc, S100A12 arranges into homomeric oligomers. To study structure-function relationships of the different oligomers, we stabilized dimeric, tetrameric and hexameric recombinant S100A12 with a chemical crosslinker and separated the complexes on a size-exclusion chromatography column. Finally, to analyze functionality and biological activity of the purified protein and its ion-induced oligomers, the cytokine release of S100A12 and LPS stimulated monocyte can be compared.
Various methods for purifying S100A12 have been described so far. Jackson et al.19, for example, published a protocol with purification via an anion-exchange column and a subsequent size-exclusion chromatography. Purification polishing on a size-exclusion column leads to good results, but&#8213;due to for example limited loading volumes&#8213;is less flexible in scalability. A different approach, published by Kiss et al.20, describes purification of tagged protein via Ni2+ affinity column as the first purification step, followed by enzymatic cleavage to remove the tag and further purification steps. In contrast to the aforecited studies19,20, the produced protein as described in this protocol is determined for experiments on immune cells. Therefore, remnant endotoxin contamination from bacterial culture is a challenge. Although different approaches for endotoxin removal have been described so far, there is no uniform method that works equally well for any given protein solution21,22.
In summary, our protocol combines the advantages of a tag-free expression in a bacterial system with efficient endotoxin removal and high yield of pure protein.

<table>
	<tbody>
		<tr>
			<td>pET11b vector</td>
			<td>Novagen</td>
			<td>&#160;</td>
			<td>&#160;</td>
		</tr>
		<tr>
			<td>BL21(DE3) competent E. coli</td>
			<td>New England Biolabs</td>
			<td>C2527</td>
			<td>&#160;</td>
		</tr>
		<tr>
			<td>&#160;</td>
			<td>&#160;</td>
			<td>&#160;</td>
			<td>&#160;</td>
		</tr>
		<tr>
			<td>100 x Non-essential amino acids</td>
			<td>Merck</td>
			<td>K 0293</td>
			<td></td>
		</tr>
		<tr>
			<td>25% HCl</td>
			<td>Carl Roth</td>
			<td>X897.1</td>
			<td></td>
		</tr>
		<tr>
			<td>4&#8722;20% Mini-PROTEAN TGX Protein Gels</td>
			<td>BioRad</td>
			<td>4561093</td>
			<td></td>
		</tr>
		<tr>
			<td>Ampicillin sodium salt</td>
			<td>Carl Roth</td>
			<td>HP62.1</td>
			<td></td>
		</tr>
		<tr>
			<td>BS3 (bis(sulfosuccinimidyl)suberate) - 50 mg</td>
			<td>ThermoFisher Scientific</td>
			<td>21580</td>
			<td></td>
		</tr>
		<tr>
			<td>Calciumchlorid Dihydrat</td>
			<td>Carl Roth</td>
			<td>5239.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Coomassie Briliant Blue R250 Destaining Solution</td>
			<td>BioRad</td>
			<td>1610438</td>
			<td></td>
		</tr>
		<tr>
			<td>Coomassie Briliant Blue R250 Staining Solution</td>
			<td>BioRad</td>
			<td>1610436</td>
			<td></td>
		</tr>
		<tr>
			<td>EasySep Human Monocyte Enrichment Kit</td>
			<td>Stemcell</td>
			<td>19059</td>
			<td>Magnetic negative cell isolation kit</td>
		</tr>
		<tr>
			<td>EDTA disodium salt dihydrate</td>
			<td>Carl Roth</td>
			<td>8043.1</td>
			<td></td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>Carl Roth</td>
			<td>3054.3</td>
			<td></td>
		</tr>
		<tr>
			<td>EndoLISA</td>
			<td>Hyglos</td>
			<td>609033</td>
			<td>Endotoxin detection assay</td>
		</tr>
		<tr>
			<td>Endotoxin-Free Ultra Pure Water</td>
			<td>Sigma-Aldrich</td>
			<td>TMS-011-A</td>
			<td>Ultrapure water for preparation of endotoxin-free buffers</td>
		</tr>
		<tr>
			<td>EndoTrap red</td>
			<td>Hyglos</td>
			<td>321063</td>
			<td>Endotoxin removal resin</td>
		</tr>
		<tr>
			<td>FBS (heat-inactivated)</td>
			<td>Gibco</td>
			<td>10270</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS, no calcium, no magnesium</td>
			<td>ThermoFisher Scientific</td>
			<td>14175053</td>
			<td></td>
		</tr>
		<tr>
			<td>Hepes</td>
			<td>Carl Roth</td>
			<td>9105</td>
			<td></td>
		</tr>
		<tr>
			<td>Hepes (high quality, endotoxin testet)</td>
			<td>Sigma-Aldrich</td>
			<td>H4034</td>
			<td></td>
		</tr>
		<tr>
			<td>hTNF-alpha - OptEia ELISA Set</td>
			<td>BD</td>
			<td>555212</td>
			<td></td>
		</tr>
		<tr>
			<td>IPTG (isopropyl-&#223;-D-thiogalactopyranosid)</td>
			<td>Carl Roth</td>
			<td>CN08.1</td>
			<td></td>
		</tr>
		<tr>
			<td>L-Glutamine (200 mM)</td>
			<td>Merck</td>
			<td>K 0282</td>
			<td></td>
		</tr>
		<tr>
			<td>LB-Medium</td>
			<td>Carl Roth</td>
			<td>X968.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Lipopolysaccharides from E. coli O55:B5</td>
			<td>Merck</td>
			<td>L6529</td>
			<td></td>
		</tr>
		<tr>
			<td>Pancoll, human</td>
			<td>PAN Biotech</td>
			<td>P04-60500</td>
			<td>Separation solution (density gradient centrifugation)</td>
		</tr>
		<tr>
			<td>Penicillin/Streptomycin (10.000 U/ml)</td>
			<td>Merck</td>
			<td>A 2212</td>
			<td></td>
		</tr>
		<tr>
			<td>Phenyl Sepharose High Performance</td>
			<td>GE Healthcare</td>
			<td>17-1082-01</td>
			<td>Resin for hydrophobic interaction chromatography</td>
		</tr>
		<tr>
			<td>Polymyxin B</td>
			<td>Invivogen</td>
			<td>tlrl-pmb</td>
			<td></td>
		</tr>
		<tr>
			<td>Protease inhibitor tablets</td>
			<td>Roche</td>
			<td>11873580001</td>
			<td></td>
		</tr>
		<tr>
			<td>Q Sepharose Fast Flow</td>
			<td>GE Healthcare</td>
			<td>17-0510-01</td>
			<td>Resin for anion-exchange chromatography</td>
		</tr>
		<tr>
			<td>RoboSep buffer</td>
			<td>Stemcell</td>
			<td>20104</td>
			<td>Cell separation buffer (section 5.1.4)</td>
		</tr>
		<tr>
			<td>RPMI 1640 Medium</td>
			<td>Merck</td>
			<td>F 1215</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride (NaCl)</td>
			<td>Carl Roth</td>
			<td>3957.2</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium hydroxide</td>
			<td>Carl Roth</td>
			<td>P031.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris Base</td>
			<td>Carl Roth</td>
			<td>4855.3</td>
			<td></td>
		</tr>
		<tr>
			<td>Zinc chloride</td>
			<td>Carl Roth</td>
			<td>T887</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Labware</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>0,45 &#181;m syringe filter</td>
			<td>Merck</td>
			<td>SLHA033SS</td>
			<td></td>
		</tr>
		<tr>
			<td>14 mL roundbottom tubes</td>
			<td>BD</td>
			<td>352059</td>
			<td></td>
		</tr>
		<tr>
			<td>2 L Erlenmyer flask</td>
			<td>Carl Roth</td>
			<td>LY98.1</td>
			<td></td>
		</tr>
		<tr>
			<td>24 well suspension plates</td>
			<td>Greiner</td>
			<td>662102</td>
			<td></td>
		</tr>
		<tr>
			<td>5 L measuring beaker</td>
			<td>Carl Roth</td>
			<td>CKN3.1</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL conical centrifuge tubes</td>
			<td>Corning</td>
			<td>430829</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL high-speed centrifuge tubes</td>
			<td>Eppendorf</td>
			<td>3,01,22,178</td>
			<td></td>
		</tr>
		<tr>
			<td>Amicon Ultra-15 Centrifugal Filter Unit MWCO 3 kDa</td>
			<td>Merck</td>
			<td>UFC900324</td>
			<td></td>
		</tr>
		<tr>
			<td>Amicon Ultra-15 Centrifugal Filter Unit MWCO 50 kDa</td>
			<td>Merck</td>
			<td>UFC905024</td>
			<td></td>
		</tr>
		<tr>
			<td>Culture dish (100 mm)</td>
			<td>Sarstedt</td>
			<td>83.3902</td>
			<td></td>
		</tr>
		<tr>
			<td>Dialysis Tubing Closures</td>
			<td>Spectrum</td>
			<td>132738</td>
			<td></td>
		</tr>
		<tr>
			<td>EasySep magnet &#39;The Big Easy`</td>
			<td>Stemcell</td>
			<td>18001</td>
			<td></td>
		</tr>
		<tr>
			<td>Fraction collector tubes 5 mL</td>
			<td>Greiner</td>
			<td>115101</td>
			<td></td>
		</tr>
		<tr>
			<td>Lumox film, 25 &#181;m, 305 mm x 40 m</td>
			<td>Sarstedt</td>
			<td>94,60,77,316</td>
			<td>Film for monocyte culture plates</td>
		</tr>
		<tr>
			<td>Spectra/Por Dialysis Membrane (3.5 kDa)</td>
			<td>Spectrum</td>
			<td>132724</td>
			<td></td>
		</tr>
		<tr>
			<td>Steritop filter unit</td>
			<td>Merck</td>
			<td>SCGPT01RE</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>37 &#176;C Incubator (with shaking)</td>
			<td>New Brunswick Scientific</td>
			<td>Innova 42</td>
			<td></td>
		</tr>
		<tr>
			<td>&#196;KTA purifier UPC 10</td>
			<td>GE Healthcare</td>
			<td></td>
			<td>FPLC System</td>
		</tr>
		<tr>
			<td>Fraction collector</td>
			<td>GE Healthcare</td>
			<td>Frac-920</td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge (with rotor A-4-81)</td>
			<td>Eppendorf</td>
			<td>5810R</td>
			<td></td>
		</tr>
		<tr>
			<td>Fixed angle rotor</td>
			<td>Eppendorf</td>
			<td>F-34-6-38</td>
			<td></td>
		</tr>
		<tr>
			<td>Mini Protean Tetra Cell</td>
			<td>BioRad</td>
			<td>1658000EDU</td>
			<td></td>
		</tr>
		<tr>
			<td>NanoPhotometer</td>
			<td>Implen</td>
			<td>P330</td>
			<td></td>
		</tr>
		<tr>
			<td>Sonicator</td>
			<td>Brandelin</td>
			<td>UW2070</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescence reader</td>
			<td>Tecan</td>
			<td>infinite M200PRO</td>
			<td></td>
		</tr>
		<tr>
			<td>pH meter</td>
			<td>Knick</td>
			<td>765</td>
			<td></td>
		</tr>
	</tbody>
</table>

purification of human s100a12 and its ion-induced oligomers for immune cell stimulation

In this protocol, we describe a method to purify human calcium-binding protein S100A12 and its ion-induced oligomers from Escherichia coli culture for immune cell stimulations. This protocol is based on a two-step chromatography strategy, which comprises protein pre-purification on an anion-exchange chromatography column and a subsequent polishing step on a hydrophobic-interaction column. This strategy produces S100A12 protein of high purity and yield at manageable costs. For functional assays on immune cells eventual remnant endotoxin contamination requires careful monitoring and further cleaning steps to obtain endotoxin-free protein. The majority of endotoxin contaminations can be excluded by anion-exchange chromatography. To deplete residual contaminations, this protocol describes a removal step with centrifugal filters. Depending on the available ion-strength S100A12 can arrange into different homomultimers. To investigate the relationship between structure and function, this protocol further describes ion-treatment of S100A12 protein followed by chemical crosslinking to stabilize S100A12 oligomers and their subsequent separation by size-exclusion chromatography. Finally, we describe a cell-based assay that confirms the biological activity of the purified protein and confirms LPS-free preparation.

Following pre-purification on the AIEX column (Figure 1A-C) and subsequent calcium-dependent HIC (Figure 2A,B), highly pure protein was obtained (Figure 2C). In addition, measurements of endotoxin revealed successful LPS removal. The LPS content following AIEX was measured in a 1:10 dilution above the assay detection limit, i.e., above 500 EU/mL. After the first filtration through a 50 kDa filter un...

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Purification of Human S100A12 and Its Ion-induced Oligomers for Immune Cell Stimulation

This protocol describes a purification method for recombinant tag-free calcium binding protein S100A12 and its ion-induced oligomers for human monocyte stimulation assays.

In this protocol, we describe a method to purify human calcium-binding protein S100A12 and its ion-induced oligomers from Escherichia coli culture for ...

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5.9K Views.  University Children's Hospital. This protocol describes a purification method for recombinant tag-free calcium binding protein S100A12 and its ion-induced oligomers for human monocyte stimulation assays.

Video: Purification of Human S100A12 and Its Ion-induced Oligomers for Immune Cell Stimulation

Purification of Specific Cell Population by Fluorescence Activated Cell Sorting (FACS)

Experimental and clinical studies often require highly purified cell populations.  FACS is a technique of choice to purify cell populations of known phenotype.  Other bulk methods of purification include panning, complement depletion and magnetic bead separation.  However, FACS has several advantages over other available methods.  FACS is the preferred method when very high purity of the desired population is required, when the target cell population expresses a very low level of the identifying marker or when cell populations require separation based on differential marker density.  In addition, FACS is the only available purification technique to isolate cells based on internal staining or intracellular protein expression, such as a genetically modified fluorescent protein marker.  FACS allows the purification of individual cells based on size, granularity and fluorescence.  In order to purify cells of interest, they are first stained with fluorescently-tagged monoclonal antibodies (mAb), which recognize specific surface markers on the desired cell population (1).  Negative selection of unstained cells is also possible.  FACS purification requires a flow cytometer with sorting capacity and the appropriate software.  For FACS, cells in suspension are passed as a stream in droplets with each containing a single cell in front of a laser.  The fluorescence detection system detects cells of interest based on predetermined fluorescent parameters of the cells.  The instrument applies a charge to the droplet containing a cell of interest and an electrostatic deflection system facilitates collection of the charged droplets into appropriate collection tubes (2).  The success of staining and thereby sorting depends largely on the selection of the identifying markers and the choice of mAb.  Sorting parameters can be adjusted depending on the requirement of purity and yield.  Although FACS requires specialized equipment and personnel training, it is the method of choice for isolation of highly purified cell populations.

Purification of Specific Cell Population by Fluorescence Activated Cell Sorting FACS

For many scientific studies requiring a biological and chemical analysis of cell populations the cells must be in a high state of purity. Fluorescence activated cell sorting (FACS) is a superior method in which to obtain pure cell populations.

Experimental and clinical studies often require highly purified cell populations.  FACS is a technique of choice to purify cell populations of known ...

Quantitative Assessment of Immune Cells in the Injured Spinal Cord Tissue by Flow Cytometry:  a Novel Use for a Cell Purification Method

Detection of immune cells in the injured central nervous system (CNS) using morphological or histological techniques has not always provided true quantitative analysis of cellular inflammation. Flow cytometry is a quick alternative method to quantify immune cells in the injured brain or spinal cord tissue. Historically, flow cytometry has been used to quantify immune cells collected from blood or dissociated spleen or thymus, and only a few studies have attempted to quantify immune cells in the injured spinal cord by flow cytometry using fresh dissociated cord tissue. However, the dissociated spinal cord tissue is concentrated with myelin debris that can be mistaken for cells and reduce cell count reliability obtained by the flow cytometer. We have advanced a cell preparation method using the OptiPrep gradient system to effectively separate lipid/myelin debris from cells, providing sensitive and reliable quantifications of cellular inflammation in the injured spinal cord by flow cytometry. As described in our recent study (Beck &amp; Nguyen et al., Brain. 2010 Feb; 133 (Pt 2): 433-47), the OptiPrep cell preparation had increased sensitivity to detect cellular inflammation in the injured spinal cord, with counts of specific cell types correlating with injury severity. Critically, novel usage of this method provided the first characterization of acute and chronic cellular inflammation after SCI to include a complete time course for polymorphonuclear leukocytes (PMNs, neutrophils), macrophages/microglia, and T-cells over a period ranging from 2 hours to 180 days post-injury (dpi), identifying a surprising novel second phase of cellular inflammation. Thorough characterization of cellular inflammation using this method may provide a better understanding of neuroinflammation in the injured CNS, and reveal an important multiphasic component of neuroinflammation that may be critical for the design and implementation of rational therapeutic treatment strategies, including both cell-based and pharmacological interventions for SCI.

Quantification of cellular inflammation in the injured/pathological CNS by flow cytometry is complicated by lipid/myelin debris that can have similar size and granulation to cells, decreasing sensitivity/accuracy. We have advanced a cell preparation method to remove myelin debris and improve cell detection by flow cytometry in the injured spinal cord.

Detection of immune cells in the injured central nervous system (CNS) using morphological or histological techniques has not always provided true ...

Flow Cytometry Analysis of Immune Cells Within Murine Aortas

Atherosclerosis is a chronic inflammatory process of medium and large size vessels that is characterized by the formation of plaques consisting of foam cells, immune cells, vascular endothelial and smooth muscle cells, platelets, extracellular matrix, and a lipid-rich core with extensive necrosis and fibrosis of surrounding tissues.1 The innate and adaptive arms of the immune response are involved in the initiation, development and persistence of atherosclerosis.2, 3 There is a significant body of evidence that different subsets of the immune cells, such as macrophages, dendritic cells, T and B lymphocytes, are present within the aortas of healthy and atherosclerosis-prone mice4. Additionally, immune cells are found in the surrounding aortic adventitia which suggests an important role of this tissue in atherogenesis.2
For some time, the quantitative detection of different types of immune cells, their activation status, and the cellular composition within the aortic wall was limited by RT-PCR and immunohistochemical methods for the study of atherosclerosis. Few attempts were made to perform flow cytometry using human aortas, and a number of problems, such as a high autofluorescence, have been reported5,6. Human atherosclerotic plaques were digested with collagenase 1, and free cells were collected and stained for CD14+/CD11c+ to highlight macrophage-derived foam cells. In this study, a "mock" channel was used to avoid false-positive staining.6 Necrotic materials accumulating during the digestion process give rise in a large amount of debris that generates a high autofluorescence in aortic samples. To resolve this problem, a panel of negative and positive controls has been proposed, but only double staining could be applied in these samples. We have developed a new flow cytometry-based method7 to analyze the immune cell composition and characterize the activation, proliferation, differentiation of immune cells in healthy and atherosclerosis-prone aorta. This method allows the investigation of the immune cell composition of the aortic wall and opens possibilities to use a broad spectrum of immunological methods for investigations of immune aspects of this disease.

This paper presents a flow cytometry-based method to investigate the immune composition of aortas. The paper also illustrates an additional technique that allows examining surrounding adventitia and vessel wall separately. This method opens possibilities to perform phenotypical analyses of aortic leukocytes and apply several immunological assays for atherosclerosis studies.

Atherosclerosis is a chronic inflammatory process of medium and large size vessels that is characterized by the formation of plaques consisting of foam ...

Isolation of Functional Cardiac Immune Cells

Cardiac immune cells are gaining interest for the roles they play in the pathological remodeling in many cardiac diseases.1-5 These immune cells, which include mast cells, T-cells and macrophages; store and release a variety of biologically active mediators including cytokines and proteases such as tryptase.6-8 These mediators have been shown to be key players in extracellular matrix metabolism by activating matrix metalloproteinases or causing collagen accumulation by modulating the cardiac fibroblasts' function.9-11 However, available techniques for isolating cardiac immune cells have been problematic because they use bacterial collagenase to digest the myocardial tissue. This technique causes activation of the immune cells and thus a loss of function. For example, cardiac mast cells become significantly less responsive to compounds that cause degranulation.12 Therefore, we developed a technique that allows for the isolation of functional cardiac immune cells which would lead to a better understanding of the role of these cells in cardiac disease.13, 14
This method requires a familiarity with the anatomical location of the rat's xiphoid process, axilla and falciform ligament, and pericardium of the heart. These landmarks are important to increase success of the procedure and to ensure a higher yield of cardiac immune cells. These isolated cardiac immune cells can then be used for characterization of functionality, phenotype, maturity, and co-culture experiments with other cardiac cells to gain a better understanding of their interactions.

This method for isolating functional immune cells from the heart provides an alternative to the conventional methods of collagenase digestion, which causes unwanted immune cell activation, resulting in a decreased responsiveness of these cells. Our method of isolation yields functional cardiac immune cells by avoiding problems associated with enzymatic digestion.

Cardiac immune cells are gaining interest for the roles they play in the pathological remodeling in many cardiac diseases.1-5  These immune cells, which ...

Human In Vitro Suppression as Screening Tool for the Recognition of an Early State of Immune Imbalance

Regulatory T cells (Tregs) are critical mediators of immune tolerance to self-antigens. In addition, they are crucial regulators of the immune response following an infection. Despite efforts to identify unique surface marker on Tregs, the only unique feature is their ability to suppress the proliferation and function of effector T cells. While it is clear that only in vitro assays can be used in assessing human Treg function, this becomes problematic when assessing the results from cross-sectional studies where healthy cells and cells isolated from subjects with autoimmune diseases (like Type 1 Diabetes-T1D) need to be compared. There is a great variability among laboratories in the number and type of responder T cells, nature and strength of stimulation, Treg:responder ratios and the number and type of antigen-presenting cells (APC) used in human in vitro suppression assays. This variability makes comparison between studies measuring Treg function difficult. The Treg field needs a standardized suppression assay that will work well with both healthy subjects and those with autoimmune diseases. We have developed an in vitro suppression assay that shows very little intra-assay variability in the stimulation of T cells isolated from healthy volunteers compared to subjects with underlying autoimmune destruction of pancreatic &beta;-cells. The main goal of this piece is to describe an in vitro human suppression assay that allows comparison between different subject groups. Additionally, this assay has the potential to delineate a small loss in nTreg function and anticipate further loss in the future, thus identifying subjects who could benefit from preventive immunomodulatory therapy1. Below, we provide thorough description of the steps involved in this procedure. We hope to contribute to the standardization of the in vitro suppression assay used to measure Treg function. In addition, we offer this assay as a tool to recognize an early state of immune imbalance and a potential functional biomarker for T1D.

Human In Vitro Suppression as Screening Tool for the Recognition of an Early State of Immune Imbalance

Tregs are potent suppressors of the immune system. There is a lack of unique surface markers to define them, hence, definitions of Tregs are primarily functional. Here we describe an optimized in vitro assay capable of identifying immune imbalance in subjects at risk to develop T1D.

Regulatory T cells (Tregs) are critical mediators of immune tolerance to self-antigens. In addition, they are crucial regulators of the immune response ...

Generation of Lymph Node-fat Pad Chimeras for the Study of Lymph Node Stromal Cell Origin

The stroma is a key component of the lymph node structure and function. However, little is known about its origin, exact cellular composition and the mechanisms governing its formation. Lymph nodes are always encapsulated in adipose tissue and we recently demonstrated the importance of this relation for the formation of lymph node stroma. Adipocyte precursor cells migrate into the lymph node during its development and upon engagement of the Lymphotoxin-b receptor switch off adipogenesis and differentiate into lymphoid stromal cells (B&#233;n&#233;zech&#160;et al.14). Based on the lymphoid stroma potential of adipose tissue, we present a method using a lymph node/fat pad chimera that allows the lineage tracing of lymph node stromal cell precursors. We show how to isolate newborn lymph nodes and EYFP+ embryonic adipose tissue and make a LN/ EYFP+ fat pad chimera. After transfer under the kidney capsule of a host mouse, the lymph node incorporates local adipose tissue precursor cells and finishes its formation. Progeny analysis of EYFP+ fat pad cells in the resulting lymph nodes can be performed by flow-cytometric analysis of enzymatically digested lymph nodes or by immunofluorescence analysis of lymph nodes cryosections. By using fat pads from different knockout mouse models, this method will provide an efficient way of analyzing the origin of the different lymph node stromal cell populations.

Generation of lymph node/fat pad chimeras for the study of lymph node stromal cell origin is described. The method involves the isolation of lymph nodes from newborn mice and embryonic fat pads, the generation of chimeric lymph node-fat pads, and their transfer under the kidney capsule of a host mouse.

The stroma is a key component of the lymph node structure and function. However, little is known about its origin, exact cellular composition and the ...

Detection of Fluorescent Nanoparticle Interactions with Primary Immune Cell Subpopulations by Flow Cytometry

Engineered nanoparticles are endowed with very promising properties for therapeutic and diagnostic purposes. This work describes a fast and reliable method of analysis by flow cytometry to study nanoparticle interaction with immune cells. Primary immune cells can be easily purified from human or mouse tissues by antibody-mediated magnetic isolation. In the first instance, the different cell populations running in a flow cytometer can be distinguished by the forward-scattered light (FSC), which is proportional to cell size, and the side-scattered light (SSC), related to cell internal complexity. Furthermore, fluorescently labeled antibodies against specific cell surface receptors permit the identification of several subpopulations within the same sample. Often, all these features vary when cells are boosted by external stimuli that change their physiological and morphological state. Here, 50 nm FITC-SiO2 nanoparticles are used as a model to identify the internalization of nanostructured materials in human blood immune cells. The cell fluorescence and side-scattered light increase after incubation with nanoparticles allowed us to define time and concentration dependence of nanoparticle-cell interaction. Moreover, such protocol can be extended to investigate Rhodamine-SiO2 nanoparticle interaction with primary microglia, the central nervous system resident immune cells, isolated from mutant mice that specifically express the Green Fluorescent Protein (GFP) in the monocyte/macrophage lineage. Finally, flow cytometry data related to nanoparticle internalization into the cells have been confirmed by confocal microscopy.

Analysis of nanoparticle interaction with defined subpopulations of immune cells by flow cytometry.

Engineered nanoparticles are endowed with very promising properties for therapeutic and diagnostic purposes. This work describes a fast and reliable ...

A Semi-automated Approach to Preparing Antibody Cocktails for Immunophenotypic Analysis of Human Peripheral Blood

Immunophenotyping of peripheral blood by flow cytometry determines changes in the frequency and activation status of peripheral leukocytes during disease and treatment. It has the potential to predict therapeutic efficacy and identify novel therapeutic targets. Whole blood staining utilizes unmanipulated blood, which minimizes artifacts that can occur during sample preparation. However, whole blood staining must also be done on freshly collected blood to ensure the integrity of the sample. Additionally, it is best to prepare antibody cocktails on the same day to avoid potential instability of tandem-dyes and prevent reagent interaction between brilliant violet dyes. Therefore, whole blood staining requires careful standardization to control for intra and inter-experimental variability.
Here, we report deployment of an automated liquid handler equipped with a two-dimensional (2D) barcode reader into a standard process of making antibody cocktails for flow cytometry. Antibodies were transferred into 2D barcoded tubes arranged in a 96 well format and their contents compiled in a database. The liquid handler could then locate the source antibody vials by referencing antibody names within the database. Our method eliminated tedious coordination for positioning of source antibody tubes. It provided versatility allowing the user to easily change any number of details in the antibody dispensing process such as specific antibody to use, volume, and destination by modifying the database without rewriting the scripting in the software method for each assay.
A proof of concept experiment achieved outstanding inter and intra- assay precision, demonstrated by replicate preparation of an 11-color, 17-antibody flow cytometry assay. These methodologies increased overall throughput for flow cytometry assays and facilitated daily preparation of the complex antibody cocktails required for the detailed phenotypic characterization of freshly collected anticoagulated peripheral blood.

Whole blood Immunophenotyping is indispensable for monitoring the human immune system. However, the number of reagents required and the instability of specific fluorochromes in premixed cocktails necessitates daily reagent preparation. Here we show that semi-automated preparation of staining antibody cocktail helps establish reliable immunophenotyping by minimizing variability in reagent dispensing.

Immunophenotyping of peripheral blood by flow cytometry determines changes in the frequency and activation status of peripheral leukocytes during disease ...

Isolation and Flow Cytometric Analysis of Immune Cells from the Ischemic Mouse Brain

Ischemic stroke initiates a robust inflammatory response that starts in the intravascular compartment and involves rapid activation of brain resident cells. A key mechanism of this inflammatory response is the migration of circulating immune cells to the ischemic brain facilitated by chemokine release and increased endothelial adhesion molecule expression. Brain-invading leukocytes are well-known contributing to early-stage secondary ischemic injury, but their significance for the termination of inflammation and later brain repair has only recently been noticed.
Here, a simple protocol for the efficient isolation of immune cells from the ischemic mouse brain is provided. After transcardial perfusion, brain hemispheres are dissected and mechanically dissociated. Enzymatic digestion with Liberase&#160;is followed by density gradient (such as Percoll) centrifugation to remove myelin and cell debris. One major advantage of this protocol is the single-layer density gradient procedure which does not require time-consuming preparation of gradients and can be reliably performed. The approach yields highly reproducible cell counts per brain hemisphere and allows for measuring several flow cytometry panels in one biological replicate. Phenotypic characterization and quantification of brain-invading leukocytes after experimental stroke may contribute to a better understanding of their multifaceted roles in ischemic injury and repair.

Inflammation plays a central role in the pathogenesis of ischemic stroke. Increasing evidence suggests that it acts as a double-edged sword which exacerbates early brain injury, but also contributes to later repair. This protocol describes the isolation of immune cells from the ischemic brain and their subsequent flow cytometric phenotyping.

Ischemic stroke initiates a robust inflammatory response that starts in the intravascular compartment and involves rapid activation of brain resident ...

Isolation, Characterization and Functional Examination of the Gingival Immune Cell Network

Immune cell networks in tissues play a vital role in mediating local immunity and maintaining tissue homeostasis, yet little is known of the resident immune cell populations in the oral mucosa and gingiva. We have established a technique for the isolation and study of immune cells from murine gingival tissues, an area of constant microbial exposure and a vulnerable site to a common inflammatory disease, periodontitis. Our protocol allows for a detailed phenotypic characterization of the immune cell populations resident in the gingiva, even at steady state. Our procedure also yields sufficient cells with high viability for use in functional studies, such as the assessment of cytokine secretion ex vivo. This combination of phenotypic and functional characterization of the gingival immune cell network should aid towards investigating the mechanisms involved in oral immunity and periodontal homeostasis, but will also advance our understanding of the mechanisms involved in local immunopathology.

We have established a technique for the isolation, phenotypic characterization and functional analysis of immune cells from murine gingiva.

Immune cell networks in tissues play a vital role in mediating local immunity and maintaining tissue homeostasis, yet little is known of the resident ...