Name: fluorescence-based monitoring of pad4 activity via a pro-fluorescence substrate analog
Uploaded: 2014-11-05T13:00:00.000+00:00
Duration: 8 min 37 s
Description: Watch this Scientific Journal Video about Fluorescence-based Monitoring of PAD4 Activity via a Pro-fluorescence Substrate Analog at JoVE.com

This work was supported by the Lehigh University Start-up Package. We thank Dr. Walter Fast for providing the GST-PAD4 plasmid for protein expression.

1. PAD4 Transformation, Expression, and Purification
<ol>
	<li>For PAD4 Transformation, transform the pGEX plasmid containing PAD4 GST fusion into chemically competent E. Coli (BL21(DE3)) cells for protein expression using the following procedure. Prepare chemically competent (calcium chloride) E. Coli (BL21(DE3)) cells according to standard protocols.
	<ol>
		<li>Thaw 50 &#956;l of previously prepared chemically competent BL21(DE3) cells on ice and mix with 1 &#956;l of the pGEX plasmid containing PAD4 gene in a 5 ml culture tube. Incubate the mixture on ice for 10 min while gently shaking every 2 min.</li>
		<li>Heat shock the cells by placing the mixture in a 42 &#176;C water bath for 40 sec. Immediately place the cell-plasmid mixture back on ice for 2 min to allow the cells to recover. Add 1 ml of sterile LB broth to the mixture and place on ice for 1 min.</li>
		<li>Incubate the heat shocked cells at 37 &#176;C, shaking at 250 rpm for 1 hr. Pipette 75 &#956;l of the transformed cells onto an ampicillin resistant agar plate and incubate at 37 &#176;C for 15 hr. Store plate at 4 &#176;C.</li>
	</ol>
	</li>
	<li>PAD4 Expression.
	<ol>
		<li>Pick 1 colony of BL21(DE3) cells from the ampicillin agar plate and place in 5 ml of LB containing 1x ampicillin. Place in incubator and shake O/N&#160;at 37 &#176;C.</li>
		<li>Transfer the 5 ml of LB (starter) into 1 L of sterile LB containing 1x ampicillin trihydrate (MW 403.45 g/mol). Place growth in a 37 &#176;C shaking incubator. Monitor the OD600 of the growth. When growth reaches an OD600 of 0.3, move growth into 16 &#176;C shaking incubator.</li>
		<li>Upon reaching an OD600 of 0.6, induce the cells with 0.3 mM isopropylthiagalactoside (IPTG, MW 238.30 g/mol). Allow cells to shake for 15 hr at 16 &#176;C.</li>
		<li>Harvest cells by centrifugation at 4,000 x g for 20 min at 0 &#176;C. Pour off supernatant and store pellet at -80 &#176;C.</li>
	</ol>
	</li>
	<li>PAD4 Purification
	<ol>
		<li>Re-suspend the pellet containing the expressed PAD4 in BL21(DE3) cells in a buffer of 50 mM NaCl (MW 58.44 g/mol), 300 mM NaH2PO4 (MW 119.98 g/mol), 10 mM Imidazole (MW 68.077 g/mol), 0.1 mM phenylmethylsulfonyl fluoride (PMSF, MW 174.94 g/mol) and 1 mM dithiothreitol (DTT, 154.25 g/mol), pH = 8.0.</li>
		<li>Lyse the cells via sonication for 15 min at 4 &#176;C. Following sonication, centrifuge the cell lysate at 20,000 x g for 20 min at 0 &#176;C. Pour off and save supernatant. Batch the supernatant with glutathione (GSH) agarose beads for 30 min at RT.</li>
		<li>Drain supernatant from GST beads/column via gravity. Wash beads with 4 x 25 ml of 1x PBS (phosphate buffered saline, pH = 8.0) at RT.</li>
		<li>Elute PAD4 with 2 x 10 ml Elution buffer, 50 mM tris (hydroxymethyl) aminomethane (Tris Base, MW 121.14 g/mol), 10 mM glutathione (GSH, MW 307.32 g/mol), pH = 8.0.</li>
		<li>Concentrate PAD4 using 100k MW cut-off centrifuge tubes and centrifuge at 4,000 x g for 20 min at 4 &#176;C. Aliquot protein into 200 &#956;l volumes in 1.0 ml microcentrifuge tubes and store at -80 &#176;C.</li>
	</ol>
	</li>
</ol>
2. Preparing Stock Solutions for Buffers
<ol>
	<li>Weigh out sodium chloride (NaCl, MW 58.44 g/mol) and prepare a 2 M solution. Mix solution until clear.</li>
	<li>Weigh out Tris(hydroxymethyl)aminomethane (Tris Base, MW 121.14 g/mol) and prepare a 2 M solution, pH = 8.0. Mix solution until clear.</li>
	<li>Weigh out calcium chloride dihydrate (CaCl2 2H2O, MW 147.01 g/mol) and prepare a 500 mM solution. Mix solution until clear.</li>
	<li>Weigh out Tris(2-carboxyethyl)phosphine (TCEP, MW 250.19 g/mol) and prepare a 200 mM solution. Mix solution until clear and store at -20 &#176;C.</li>
	<li>Weigh out dithiothreitol (DTT, MW 154.25 g/mol) and prepare a 1 M solution. Mix solution until clear and store at -20 &#176;C.</li>
	<li>Prepare a 0.5% solution of Triton X-100.</li>
	<li>Weigh out ethylenediaminetetraacetic acid (EDTA, MW 292.24 g/mol) and prepare a 100 mM solution. Mix solution until clear.</li>
	<li>Weigh out Z-&#8203;Arg-&#8203;Arg-&#8203;7-&#8203;amido-&#8203;4-&#8203;methylcoumarin hydrochloride (ZRcoum, MW 621.69 g/mol) and prepare a 10 mM solution in dimethyl sulfoxide (DMSO).</li>
</ol>
3. PAD4 Assay at 37 &#176;C
<ol>
	<li>From 10 mM ZRcoum stock, prepare a 125 &#956;M solution of ZRcoum in water. This 125 &#956;M ZRcoum solution will be Solution A.</li>
	<li>Prepare a buffer of 62.5 mM NaCl, 62.5 mM Tris, 12.5 mM CaCl2, 6.25 mM DTT, and 5 &#956;M PAD4 (pH = 8.0). This will be Solution B.</li>
	<li>Prepare a buffer of 62.5 mM NaCl, 62.5 mM Tris, 12.5 mM CaCl2, and 6.25 mM DTT (pH = 8.0). This will be Solution C.</li>
	<li>Weigh out Trypsin, crystalline (from bovine pancreas) and prepare a 10 mg/ml in 100 mM EDTA. Mix solution until clear and store at -20 &#176;C. This will be Solution D.</li>
	<li>Obtain a 96-well plate and designate a number of wells for controls (no PAD4) and test wells (+ PAD4).</li>
	<li>Pipette 160 &#956;l of Solution B into test wells and pipette 160 &#956;l of Solution C into control wells. Incubate for 15 min at 37 &#176;C.</li>
	<li>Pipette 40 &#956;l of Solution A into all wells and incubate at 37 &#176;C for 45 min.</li>
	<li>Pipette 10 &#956;l of dH2O to half of the control wells and half of the test wells. To the other half of control wells and test controls, pipette 10 &#956;l of Solution D. Incubate for 10 min at 37 &#176;C.</li>
	<li>Obtain a fluorescence reading on a multimodal reader with an excitation wavelength of 345 nm and an emission wavelength of 465 nm.</li>
</ol>
4. PAD4 Assay at RT
<ol>
	<li>From 10 mM ZRcoum stock, prepare a 50 &#956;M solution of ZRcoum in water. This 50 &#956;M ZRcoum solution will be Solution A.</li>
	<li>Prepare a buffer of 100 mM NaCl, 100 mM Tris, 20 mM CaCl2, 2 mM TCEP, 0.02% Triton X-100, and 8 &#956;M PAD4 (pH = 8.0). This will be Solution B.</li>
	<li>Prepare a buffer of 100 mM NaCl, 100 mM Tris, 20 mM CaCl2, 2 mM TCEP, and 0.02% Triton X-100 (pH = 8.0). This will be Solution C.</li>
	<li>Weigh out Trypsin, crystalline (from bovine pancreas) and prepare a 10 mg/ml in 100 mM EDTA. Mix solution until clear and store at -20 &#176;C. This will be Solution D.</li>
	<li>Obtain a 384-well plate and designate wells for controls (no PAD4) and test wells (+ PAD4).</li>
	<li>Pipette 25 &#956;l of Solution B into test wells and pipette 25 &#956;l of Solution C into control wells. Allow buffer with/without PAD4 to incubate in wells for 20 min at RT.</li>
	<li>Pipette 25 &#956;l of Solution C into all wells and incubate for 45 min at RT.</li>
	<li>Pipette 10 &#956;l of dH2O to half of the control wells and half of the test wells. To the other half of control wells and test controls, pipette 10 &#956;l of Solution D. Incubate at RT for 10 min.</li>
	<li>Obtain a fluorescence reading on a multimodal reader with an excitation wavelength of 345 nm and an emission wavelength of 465 nm.</li>
</ol>
5. PAD4 Time Course Assays at 37 &#176;C and RT
<ol>
	<li>From 10 mM ZRcoum stock, prepare a 125 &#956;M solution of ZRcoum in water. This 125 &#956;M ZRcoum solution will be Solution A.</li>
	<li>Prepare a buffer of 62.5 mM NaCl, 62.5 mM Tris, 12.5 mM CaCl2, 6.25 mM DTT, and 5 &#956;M PAD4 (pH = 8.0). This will be Solution B.</li>
	<li>Weigh out Trypsin, crystalline (from bovine pancreas) and prepare a 10 mg/ml in 100 mM EDTA. Mix solution until clear and store at -20 &#176;C. This will be Solution C.</li>
	<li>Obtain 2 96-well plates and pipet 160 &#956;l of Solution B into # amount of wells (# of wells will depend on how many time points).</li>
	<li>Incubate one 96-well plate at 37 &#176;C. Incubate the other 96 well plate at RT.</li>
	<li>Pipette 40 &#956;l of Solution A into all wells in both 96-well plates and immediately pipette 10 &#956;l of Solution C into the t = 0 min wells. Continue adding 10 &#956;l of Solution C at specific time points until all wells have been quenched.</li>
	<li>Obtain a fluorescence reading on a multimodal reader with an excitation wavelength of 345 nm and an emission wavelength of 465 nm.</li>
</ol>
6. PAD4 Inhibition Assay at RT
<ol>
	<li>From 10 mM ZRcoum stock, prepare a 83 &#956;M solution of ZRcoum in water. This 83 &#956;M ZRcoum solution will be Solution A.</li>
	<li>Weigh out Cl-amidine (MW 424.8 g/mol) and prepare a 100 &#956;M stock solution. This will be Solution B.</li>
	<li>Prepare a buffer of 100 mM NaCl, 100 mM Tris, 20 mM CaCl2, 2 mM TCEP, 0.02% Triton X-100, and 8 &#956;M PAD4 (pH = 8.0). This will be Solution C.</li>
	<li>Prepare a buffer of 100 mM NaCl, 100 mM Tris, 20 mM CaCl2, 2 mM TCEP, and 0.02% Triton X-100 (pH = 8.0). This will be Solution D.</li>
	<li>Weigh out Trypsin, crystalline (from bovine pancreas) and prepare a 10 mg/ml in 100 mM EDTA. Mix solution until clear and store at -20 &#176;C. This will be Solution E.</li>
	<li>Obtain a 384-well plate and designate wells for controls (no PAD4), test wells (+ PAD4), and inhibition wells (PAD4 + Cl-amidine).</li>
	<li>Pipette 25 &#956;l of Solution C into test wells and pipette 25 &#956;l of Solution D into control wells. Allow buffer with/without PAD4 to incubate in wells for 20 min at RT.</li>
	<li>Pipette 10 &#956;l of Solution B into inhibition wells and incubate for 20 min at RTe.</li>
	<li>Pipette 15 &#956;l of Solution A into all wells and incubate for 45 min at RT.</li>
	<li>Pipette 10 &#956;l of Solution E to all wells. Incubate at RT for 10 min.</li>
	<li>Obtain a fluorescence reading on a multimodal reader with an excitation wavelength of 345 nm and an emission wavelength of 465 nm.</li>
</ol>
7. Crude Cell Lysate Assay at RT
<ol>
	<li>After harvesting cells from PAD4 1 L growth, resuspend cells in 10 ml of 50 mM Tris, 50 mM NaCl, 0.1 mM phenylmethylsulfonyl fluoride (PMSF, MW 174.94 g/mol) and 1 mM dithiothreitol (DTT, 154.25 g/mol), pH = 8.0.</li>
	<li>Lyse the cells via sonication for 15 min at 4 &#176;C. Centrifuge the cell lysate at 20,000 x g for 20 min at 0 &#176;C. Concentrate the clear supernatant to half the volume by centrifugation at 4,000 rpm at 4 &#176;C using 100k MW cut-off centrifuge tubes.</li>
	<li>From 10 mM ZRcoum stock, prepare a 250 &#956;M solution of ZRcoum in water. This 250 &#956;M ZRcoum solution will be Solution A.</li>
	<li>Prepare a buffer of 100 mM NaCl, 100 mM Tris, 20 mM CaCl2, and 2 mM TCEP (pH = 8.0). This will be Solution B.</li>
	<li>Weigh out Trypsin, crystalline (from bovine pancreas) and prepare a 10 mg/ml in 100 mM EDTA. Mix solution until clear and store at -20 &#176;C. This will be Solution C.</li>
	<li>Obtain a black 384-well plate and designate wells for controls (no cell lysate) and test wells (+ cell lysate).</li>
	<li>Pipette 100 &#956;l of Solution B into all wells and pipette 60 &#956;l of cell lysate into test wells. Incubate for 20 min at RT.</li>
	<li>Pipette 40 &#956;l of Solution A into all wells and incubate for 45 min at RT.</li>
	<li>Pipette 10 &#956;l of Solution C to all wells. Incubate at RT for 10 min.</li>
	<li>Obtain a fluorescence reading on multimodal reader with an excitation wavelength of 345 nm and an emission wavelength of 465 nm.</li>
</ol>

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R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein+arginine+deiminase+4:+a+target+for+an+epigenetic+cancer+therapy.">Protein arginine deiminase 4: a target for an epigenetic cancer therapy.</a> Cell. Mol. Life Sci. 68, 709-720 (2011).</li><li>Luo, Y., Knuckley, B., Lee, Y. H., Stallcup, M. R., Thompson, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+fluoroacetamidine-based+inactivator+of+protein+arginine+deiminase+4:+design,+synthesis,+and+in+vitro+and+in+vivo+evaluation.">A fluoroacetamidine-based inactivator of protein arginine deiminase 4: design, synthesis, and in vitro and in vivo evaluation.</a> J. Am. Chem. Soc. 128, 1092-1093 (2006).</li><li>Knuckley, 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=Substrate+specificity+and+kinetic+studies+of+PADs.">Substrate specificity and kinetic studies of PADs.</a> Biochemistry. 1, 4852-4863 (2010).</li><li>Knipp, M., Vasak, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+colorimetric+96-well+microtiter+plate+assay+for+the+determination+of+enzymatically+formed+citrulline.">A colorimetric 96-well microtiter plate assay for the determination of enzymatically formed citrulline.</a> Anal. Biochem. 286, 257-264 (2000).</li><li>Knuckley, 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=A+fluopol-ABPP+HTS+assay+to+identify+PAD+inhibitors.">A fluopol-ABPP HTS assay to identify PAD inhibitors.</a> Chem. Commun. 46, 7175-7177 (2010).</li><li>Bicker, K. L., Subramanian, V., Chumanevich, A. A., Hofseth, L. J., Thompson, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Seeing+citrulline:+development+of+a+phenylglyoxal-based+probe+to+visualize+protein+citrullination.">Seeing citrulline: development of a phenylglyoxal-based probe to visualize protein citrullination.</a> J. Am. Chem. Soc. 134, 17015-17018 (2012).</li><li>Wang, Q., Priestman, M. A., Lawrence, D. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Monitoring+of+protein+arginine+deiminase+activity+by+using+fluorescence+quenching:+multicolor+visualization+of+citrullination.">Monitoring of protein arginine deiminase activity by using fluorescence quenching: multicolor visualization of citrullination.</a> Angew. Chem. Int. Ed. Engl. 52, 2323-2325 (2013).</li><li>Jones, J. E., 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=Synthesis+and+screening+of+a+haloacetamidine+containing+library+to+identify+PAD4+selective+inhibitors.">Synthesis and screening of a haloacetamidine containing library to identify PAD4 selective inhibitors.</a> ACS Chem. Biol. 7, 160-165 (2012).</li><li>Dreyton, C. J., 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=.">.</a> Optimization and characterization of a pan protein arginine deiminase (PAD) inhibitor. , (2010).</li><li>Shimoyama, S., 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=Deimination+stabilizes+histone+H2A/H2B+dimers+as+revealed+by+electrospray+ionization+mass+spectrometry.">Deimination stabilizes histone H2A/H2B dimers as revealed by electrospray ionization mass spectrometry.</a> J. Mass Spectrom. 45, 900-908 (2010).</li><li>Luo, Y., 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=Inhibitors+and+inactivators+of+protein+arginine+deiminase+4:+functional+and+structural+characterization.">Inhibitors and inactivators of protein arginine deiminase 4: functional and structural characterization.</a> Biochemistry. 45, 11727-11736 (2006).</li></ol>

The authors have nothing to disclose.

Herein, a fluorescence based assay was successfully developed that monitors the activity of PAD4. The assay has proven to be incredibly robust in a number of high-throughput conditions and is also compatible with whole cell lysates37.
The affinity between the histone proteins and the DNA strands surrounding it loosens due the neutralization of the positive charge on the arginine side chain upon citrullination. It was envisioned that the neutralization of the arginine side-chain coul...

A large number of mammalian proteins are heavily modified by the action of enzymes following the biosynthesis of proteins by the ribosome. These post-translational modifications (PTMs) can greatly increase the functional diversity of the proteome by changing the size, charge, structure, and oligomerization state (amongst other features) of proteins1-3. As a result, the change in protein structure can lead to physiological consequences, such as protein degradation, cellular differentiation, signaling, modulation in gene expression, and protein-protein interactions. While these modifications are prevalent in a large percentage of all human proteins, the terminal ends on histone proteins undergo an unusually high number of covalent modifications4. Histone proteins are a family of structural proteins that facilitate the condensation of genomic DNA. Covalent modifications of the unstructured histone tails are carried out and regulated by a series of enzymes that can catalyze the covalent modification of residues (writers), reverse the same modifications (erasers), and distinguish among the changes being imprinted onto the histone tails (readers)5-7. In fact, most of the known PTMs can be observed within this short segment of the histone including methylation, phosphorylation, acetylation, sumoylation, ubiquitination, and citrullination8.
Citrullination involves the conversion of peptidyl-arginine to the non-tRNA encoded peptidyl-citrulline (Figure 1A). The proteins, responsible for this side chain neutralization, are members of the PAD (peptide arginine deiminase) protein family, all of which are calcium-dependent enzymes.9,10. To date, five members of the PAD family have been described (PAD1, PAD2, PAD3, PAD4, and PAD6). Each member of this family appears to target distinct cellular proteins and also displays unique tissue distribution profiles. PAD4 is the only member of this protein family known to be localized within the nucleus via a nuclear localization sequence11. Accordingly, it has been shown to deiminate a number of nuclear targets, including arginine side chains on the N-terminal tails of histones H2A arginine residue 3 (H2R3), H3 (H3R2, H3R17, and H3R26) and H4 (H4R3)12,13. While each of the PAD isozymes have specific and critical physiological functions, PAD4 has received considerably more attention due to its role in a number of human processes in both diseased and healthy cells. Recently, PAD4 was shown to be a member of the pluripotency transcriptional network14. Both PAD4 expression levels and activity were shown to be elevated during reprogramming and ground-state pluripotent states in mice. By controlling the regulation of stem-cell genes, PAD4 may retain a pivotal role in cellular reprogramming efficiency. PAD4 has also been implicated in the formation of neutrophil extracellular traps, which upon binding of pathogens enable their system clearance. The hypercitrullination of histone proteins by PAD4 induces decondensation of the chromatin, which serves as the base material for the encapsulation of pathogenic bacteria by the extracellular trap, thus warding off bacterial infections15,16.
Additionally, PAD4 has been found to play an active role in a number of human diseases. It has been previously shown that aberrant expression of PAD4 is associated with the onset and severity of rheumatoid arthritis, Alzheimer&#8217;s, Parkinson&#8217;s disease, and multiple sclerosis17. In fact, the presence of anti-citrullinated protein antibodies is one of the most reliable and definitive diagnostic and prognostic biomarkers of rheumatoid arthritis18. Likewise, dysregulated PAD4 activity has recently been observed in a number of human cancers including ovarian, breast, lung, and esophageal cancers19-22. The link between PAD4 and cancer has been shown to be mediated through the ELK1 oncogene or via the p53 tumor suppressor protein22,23 and previous work has suggested that PAD4 could be a novel anti-cancer therapeutic target17,24,25. As a proof of principle study, the depletion of PAD4 via shRNA in the colorectal cancer cell line HCT116 was revealed to be sufficient for inducing apoptosis and cell cycle arrest26. A recently developed irreversible PAD4 inhibitor led to a seventy percent reduction in tumor mass in mice27. Quite remarkably, PAD4 inhibition appeared to act as a targeted therapy that resulted in selective killing of cancerous cells while sparing untransformed cells.
The use of small molecules to turn off the function of PAD4 may prove to be a powerful new strategy to target cancer cells or to augment existing cancer chemotherapeutics28. Unfortunately, a potent reversible PAD4 inhibitor has yet to be discovered. A number of covalent inhibitors have been developed using the chloro/fluoro imidine handle that mimics the arginine substrate27,29,30 and have proven to be practical tools in understanding the role of PAD4 in both healthy and diseased state cells. However, these molecules inhibit all of the active PADs with similar potency. Therefore, the need for a facile assay that reports on the activity of PAD4 is crucial. To date, PAD4 assays have been described that link the release of ammonia from the reaction to a colorimetric readout31, utilize a fluorescently labeled chloroamidine substrate analog for fluorescence polarization assay32, rely on the acid-assisted reaction between glyoxal and citrulline33, and couple the PAD4 activity to a fluorescence dequenching step34. Of these, only the covalent modifier haloacetamidine strategy has proven to be compatible with high-throughput screening platforms32,35,36. We describe a facile fluorescence based assay that reliably measures the activity of PAD4. The assay, which displays a strong signal-to-noise ratio, speed of analysis, and robustness of measurement, has the potential to discover a truly potent and selective PAD4 inhibitor.

<table><tbody><tr><td>LB Broth, Miller (LURIA-BERTANI)</td><td>Amresco</td><td>J106-2KG</td><td>&lt;a href=&quot;http://www.amresco-inc.com/&quot;&gt;http://www.amresco-inc.com&lt;/a&gt;</td></tr><tr><td>Ampicillin Trihydrate (Off-white Powder), Fisher BioReagents</td><td>Fisher Scientific</td><td>BP902-25&amp;nbsp;</td><td>&lt;a href=&quot;http://www.fishersci.com/&quot;&gt;http://www.fishersci.com&lt;/a&gt;</td></tr><tr><td>Isopropylthiagalactonse (IPTG)</td><td>Life Technogolies</td><td>15529-019</td><td>&lt;a href=&quot;http://www.lifetechnologies.com/&quot;&gt;http://www.lifetechnologies.com&lt;/a&gt;</td></tr><tr><td>All Buffer Salts&amp;nbsp;</td><td>Fisher Scientific</td><td></td><td>Can purchase from Fisher Scieintifc; VWR; Acros; Sigma-Aldrich; etc.</td></tr><tr><td>Protino Glutathione (GSH) agarose&amp;nbsp;</td><td>Machery-Nagel</td><td>745500.10</td><td>Store at 4 &amp;deg;C; http://www.mn-net.com/</td></tr><tr><td>Chloroamidine&amp;nbsp;</td><td>Cayman Chemical&amp;nbsp;</td><td>10599</td><td>Store at -20 &amp;deg;C; https://www.caymanchem.com/app/template/Home.vm</td></tr><tr><td>Triton X-100</td><td>Sigma Aldrich</td><td>X100-100ML</td><td>&lt;a href=&quot;http://www.sigmaaldrich.com/&quot;&gt;www.sigmaaldrich.com&lt;/a&gt;</td></tr><tr><td>Corning/Nunclon MicroWell plates (96 and 384)</td><td></td><td></td><td>Purchase from Corning; Sigma-Aldrich</td></tr><tr><td>Tecan Infinite 200 / Tecan i-control microplate reader software</td><td></td><td></td><td>&lt;a href=&quot;http://www.tecan.com/&quot;&gt;www.tecan.com&lt;/a&gt;</td></tr><tr><td>Europium Ex. 340/40 Em. 475/15 filter</td><td></td><td></td><td>&lt;a href=&quot;http://www.perkinelmer.com/&quot;&gt;http://www.perkinelmer.com&lt;/a&gt;</td></tr></tbody></table>

fluorescence-based monitoring of pad4 activity via a pro-fluorescence substrate analog

Post-translational modifications may lead to altered protein functional states by increasing the covalent variations on the side chains of many protein substrates. The histone tails represent one of the most heavily modified stretches within all human proteins. Peptidyl-arginine deiminase 4 (PAD4) has been shown to convert arginine residues into the non-genetically encoded citrulline residue. Few assays described to date have been operationally facile with satisfactory sensitivity. Thus, the lack of adequate assays has likely contributed to the absence of potent non-covalent PAD4 inhibitors. Herein a novel fluorescence-based assay that allows for the monitoring of PAD4 activity is described. A pro-fluorescent substrate analog was designed to link PAD4 enzymatic activity to fluorescence liberation upon the addition of the protease trypsin. It was shown that the assay is compatible with high-throughput screening conditions and has a strong signal-to-noise ratio. Furthermore, the assay can also be performed with crude cell lysates containing over-expressed PAD4.

Initially, it was demonstrated that ZRcoum can report on the activity of PAD4 in a 96-well plate format. Wells were incubated with the substrate ZRcoum and in the presence/absence of PAD4. Following an incubation period of 45 min at 37 &#176;C, the fluorescence was measured with a 340/40 &#8211; 475/15 nm filter (Figure 2A). As expected, the fluorescence levels remained low as the fluorophore within ZRcoum (or the citrullinated ZRcoum) r...

Watch this Scientific Journal Video about Fluorescence-based Monitoring of PAD4 Activity via a Pro-fluorescence Substrate Analog at JoVE.com

Fluorescence-based Monitoring of PAD4 Activity via a Pro-fluorescence Substrate Analog

PAD4 is an enzyme responsible for the conversion of peptidyl-arginine to peptidyl-citrulline. Dysregulation of PAD4 has been implicated in a number of human diseases. A facile and high-throughput compatible fluorescence based PAD4 assay is described.

Post-translational modifications may lead to altered protein functional states by increasing the covalent variations on the side chains of many protein ...

fluorescence-based-monitoring-pad4-activity-via-pro-fluorescence

Universidad de Cartagena UDEC

Research

JoVE Journal

Chemistry

9.8K Views.  Lehigh University. PAD4 is an enzyme responsible for the conversion of peptidyl-arginine to peptidyl-citrulline. Dysregulation of PAD4 has been implicated in a number of human diseases. A facile and high-throughput compatible fluorescence based PAD4 assay is described.

Video: Fluorescence-based Monitoring of PAD4 Activity via a Pro-fluorescence Substrate Analog

A Fluorescence-based Assay of Phospholipid Scramblase Activity

Scramblases translocate phospholipids across the membrane bilayer bidirectionally in an ATP-independent manner. The first scramblase to be identified and biochemically verified was opsin, the apoprotein of the photoreceptor rhodopsin. Rhodopsin is a G protein-coupled receptor localized in rod photoreceptor disc membranes of the retina where it is responsible for the perception of light. Rhodopsin&#39;s scramblase activity does not depend on its ligand 11-cis-retinal, i.e., the apoprotein opsin is also active as a scramblase. Although constitutive and regulated phospholipid scrambling play an important role in cell physiology, only a few phospholipid scramblases have been identified so far besides opsin. Here we describe a fluorescence-based assay of opsin&#39;s scramblase activity. Opsin is reconstituted into large unilamellar liposomes composed of phosphatidylcholine, phosphatidylglycerol and a trace quantity of fluorescent NBD-labeled PC (1-palmitoyl-2-{6-[7-nitro-2-1,3-benzoxadiazole-4-yl)amino]hexanoyl}-sn-glycero-3-phosphocholine). Scramblase activity is determined by measuring the extent to which NBD-PC molecules located in the inner leaflet of the vesicle are able to access the outer leaflet where their fluorescence is chemically eliminated by a reducing agent that cannot cross the membrane. The methods we describe have general applicability and can be used to identify and characterize scramblase activities of other membrane proteins.

We describe a fluorescence-based assay to measure phospholipid scrambling in large unilamellar liposomes reconstituted with opsin.

Scramblases translocate phospholipids across the membrane bilayer bidirectionally in an ATP-independent manner. The first scramblase to be identified and ...

Biochemistry

Click-Chemistry Based Fluorometric Assay for Apolipoprotein N-acyltransferase from Enzyme Characterization to High-Throughput Screening

Lipoproteins from proteobacteria are posttranslationally modified by fatty acids derived from membrane phospholipids by the action of three integral membrane enzymes, resulting in triacylated proteins. The first step in the lipoprotein modification pathway involves the transfer of a diacylglyceryl group from phosphatidylglycerol onto the prolipoprotein, resulting in diacylglyceryl prolipoprotein. In the second step, the signal peptide of prolipoprotein is cleaved, forming an apolipoprotein, which in turn is modified by a third fatty acid derived from a phospholipid. This last step is catalyzed by apolipoprotein N-acyltransferase (Lnt). The lipoprotein modification pathway is essential in most &#947;-proteobacteria, making it a potential target for the development of novel antibacterial agents. Described here is a sensitive assay for Lnt that is compatible with high-throughput screening of small inhibitory molecules. The enzyme and substrates are membrane-embedded molecules; therefore, the development of an in vitro test is not straightforward. This includes the purification of the active enzyme in the presence of detergent, the availability of alkyne-phospholipids and diacylglyceryl peptide substrates, and the reaction conditions in mixed micelles. Furthermore, in order to use the activity test in a high-throughput screening (HTS) setup, direct readout of the reaction product is preferred over coupled enzymatic reactions. In this fluorometric enzyme assay, the alkyne-triacylated peptide product is rendered fluorescent through a click-chemistry reaction and detected in a multiwell plate format. This method is applicable to other acyltransferases that use fatty acid-containing substrates, including phospholipids and acyl-CoA.

Presented here is a sensitive fluorescence assay to monitor apolipoprotein N-acyltransferase activity using diacylglyceryl peptide and alkyne-phospholipids as substrates with click-chemistry.

Lipoproteins from proteobacteria are posttranslationally modified by fatty acids derived from membrane phospholipids by the action of three integral ...

Fluorescence-Based Measurements of Phosphatidylserine/Phosphatidylinositol 4-Phosphate Exchange Between Membranes

Several members of the evolutionarily conserved oxysterol-binding protein (OSBP)-related proteins(ORP)/OSBP homologs (Osh) family have recently been found to represent a novel lipid transfer protein (LTP) group in yeast and human cells. They transfer phosphatidylserine (PS) from the endoplasmic reticulum (ER) to the plasma membrane (PM) via PS/phosphatidylinositol 4-phosphate (PI(4)P) exchange cycles. This finding allows a better understanding of how PS, which is critical for signaling processes, is distributed throughout the cell and the investigation of the link between this process and phosphoinositide (PIP) metabolism. The development of new fluorescence-based protocols has been instrumental in the discovery and characterization of this new cellular mechanism in vitro at the molecular level. This paper describes the production and the use of two fluorescently labelled lipid sensors, NBD-C2Lact and NBD-PHFAPP, to measure the ability of a protein to extract PS or PI(4)P and to transfer these lipids between artificial membranes. First, the protocol describes how to produce, label, and obtain high-purity samples of these two constructs. Secondly, this paper explains how to use these sensors with a fluorescence microplate reader to determine whether a protein can extract PS or PI(4)P from liposomes, using Osh6p as a case study. Finally, this protocol shows how to accurately measure the kinetics of PS/PI(4)P exchange between liposomes of defined lipid composition and to determine lipid transfer rates by fluorescence resonance energy transfer (FRET) using a standard fluorometer.

Here, we describe protocols using fluorescent lipid sensors and liposomes to determine whether a protein extracts and transports phosphatidylserine or phosphatidylinositol 4-phosphate in vitro.

Several members of the evolutionarily conserved oxysterol-binding protein (OSBP)-related proteins(ORP)/OSBP homologs (Osh) family have recently been found ...

An Optimized Single-Molecule Pull-Down Assay for Quantification of Protein Phosphorylation

Phosphorylation is a necessary posttranslational modification that regulates protein function and directs cell signaling outcomes. Current methods to measure protein phosphorylation cannot preserve the heterogeneity in phosphorylation across individual proteins. The single-molecule pull-down (SiMPull) assay was developed to investigate the composition of macromolecular complexes via immunoprecipitation of proteins on a glass coverslip followed by single-molecule imaging. The current technique is an adaptation of SiMPull that provides robust quantification of the phosphorylation state of full-length membrane receptors at the single-molecule level. Imaging thousands of individual receptors in this way allows for quantifying protein phosphorylation patterns. The present protocol details the optimized SiMPull procedure, from sample preparation to imaging. Optimization of glass preparation and antibody fixation protocols further enhances data quality. The current protocol provides code for the single-molecule data analysis that calculates the fraction of receptors phosphorylated within a sample. While this work focuses on phosphorylation of the epidermal growth factor receptor (EGFR), the protocol can be generalized to other membrane receptors and cytosolic signaling molecules.

The present protocol describes sample preparation and data analysis to quantify protein phosphorylation using an improved single-molecule pull-down (SiMPull) assay.

Phosphorylation is a necessary posttranslational modification that regulates protein function and directs cell signaling outcomes. Current methods to ...

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

Biology

Studying Protease-Activated Receptor Ligands in Endothelial Cells

Characterizing Modulators of Protease-Activated Receptors with a Calcium Mobilization Assay Using a Plate Reader

Changes in calcium concentration in cells are rapidly monitored in a high-throughput fashion with the use of intracellular, fluorescent, calcium-binding dyes and imaging instruments that can measure fluorescent emissions from up to 1,536 wells simultaneously. However, these instruments are much more expensive and can be challenging to maintain relative to widely available plate readers that scan wells individually. Described here is an optimized plate reader assay for use with an endothelial cell line (EA.hy926) to measure the protease-activated receptor (PAR)-driven activation of G&#945;q signaling and subsequent calcium mobilization using the calcium-binding dye Fluo-4. This assay has been used to characterize a range of PAR ligands, including the allosteric PAR1-targeting anti-inflammatory "parmodulin" ligands identified in the Dockendorff lab. This protocol obviates the need for an automated liquid handler and permits the medium-throughput screening of PAR ligands in 96-well plates and should be applicable to the study of other receptors that initiate calcium mobilization.

Author Spotlight: Developing Parmodulins to Target Protease-Activated Receptors for Inflammation Control

An improved protocol for a calcium mobilization assay with endothelial cells, used to identify ligands of protease-activated receptors (PARs), has been developed. The new protocol reduces total assay time by 90-120 min and yields reproducible concentration-response curves.

Changes in calcium concentration in cells are rapidly monitored in a high-throughput fashion with the use of intracellular, fluorescent, calcium-binding ...

FRET Microscopy for Real-time Monitoring of Signaling Events in Live Cells Using Unimolecular Biosensors

F&ouml;rster resonance energy transfer (FRET) microscopy continues to gain increasing interest as a technique for real-time monitoring of biochemical and signaling events in live cells and tissues. Compared to classical biochemical methods, this novel technology is characterized by high temporal and spatial resolution. FRET experiments use various genetically-encoded biosensors which can be expressed and imaged over time in situ or in vivo1-2. Typical biosensors can either report protein-protein interactions by measuring FRET between a fluorophore-tagged pair of proteins or conformational changes in a single protein which harbors donor and acceptor fluorophores interconnected with a binding moiety for a molecule of interest3-4. Bimolecular biosensors for protein-protein interactions include, for example, constructs designed to monitor G-protein activation in cells5, while the unimolecular sensors measuring conformational changes are widely used to image second messengers such as calcium6, cAMP7-8, inositol phosphates9 and cGMP10-11. Here we describe how to build a customized epifluorescence FRET imaging system from single commercially available components and how to control the whole setup using the Micro-Manager freeware. This simple but powerful instrument is designed for routine or more sophisticated FRET measurements in live cells. Acquired images are processed using self-written plug-ins to visualize changes in FRET ratio in real-time during any experiments before being stored in a graphics format compatible with the build-in ImageJ freeware used for subsequent data analysis. This low-cost system is characterized by high flexibility and can be successfully used to monitor various biochemical events and signaling molecules by a plethora of available FRET biosensors in live cells and tissues. As an example, we demonstrate how to use this imaging system to perform real-time monitoring of cAMP in live 293A cells upon stimulation with a &beta;-adrenergic receptor agonist and blocker.

F&#246;rster resonance energy transfer (FRET) microscopy is a powerful technique for real-time monitoring of signaling events in live cells using various biosensors as reporters. Here we describe how to build a customized epifluorescence FRET imaging system from commercially available components and how to use it for FRET experiments.

F&ouml;rster resonance energy transfer (FRET) microscopy  continues to gain increasing interest as a technique for real-time monitoring  of biochemical ...

Visualizing Protein Kinase A Activity In Head-fixed Behaving Mice Using In Vivo Two-photon Fluorescence Lifetime Imaging Microscopy

Neuromodulation exerts powerful control over brain function. Dysfunction of neuromodulatory systems results in neurological and psychiatric disorders. Despite their importance, technologies for tracking neuromodulatory events with cellular resolution are just beginning to emerge. Neuromodulators, such as dopamine, norepinephrine, acetylcholine, and serotonin, trigger intracellular signaling events via their respective G protein-coupled receptors to modulate neuronal excitability, synaptic communications, and other neuronal functions, thereby regulating information processing in the neuronal network. The above mentioned neuromodulators converge onto the cAMP/protein kinase A (PKA) pathway. Therefore, in vivo PKA imaging with single-cell resolution was developed as a readout for neuromodulatory events in a manner analogous to calcium imaging for neuronal electrical activities. Herein, a method is presented to visualize PKA activity at the level of individual neurons in the cortex of head-fixed behaving mice. To do so, an improved A-kinase activity reporter (AKAR), called tAKAR&#945;, is used, which is based on F&#246;rster resonance energy transfer (FRET). This genetically-encoded PKA sensor is introduced into the motor cortex via in utero electroporation (IUE) of DNA plasmids, or stereotaxic injection of adeno-associated virus (AAV). FRET changes are imaged using two-photon fluorescence lifetime imaging microscopy (2pFLIM), which offers advantages over ratiometric FRET measurements for quantifying FRET signal in light-scattering brain tissue. To study PKA activities during enforced locomotion, tAKAR&#945; is imaged through a chronic cranial window above the cortex of awake, head-fixed mice, which run or rest on a speed-controlled motorized treadmill. This imaging approach will be applicable to many other brain regions to study corresponding behavior-induced PKA activities and to other FLIM-based sensors for in vivo imaging.

A procedure is presented to visualize protein kinase A activities in head-fixed, behaving mice. An improved A-kinase activity reporter, tAKAR&#945;, is expressed in cortical neurons and made accessible for imaging through a cranial window. Two-photon fluorescence lifetime imaging microscopy is used to visualize PKA activities in vivo during enforced locomotion.

Neuromodulation exerts powerful control over brain function. Dysfunction of neuromodulatory systems results in neurological and psychiatric disorders. ...

Neuroscience

Utilizing pHluorin-tagged Receptors to Monitor Subcellular Localization and Trafficking

Understanding membrane protein trafficking, assembly, and expression requires an approach that differentiates between those residing in intracellular organelles and those localized on the plasma membrane. Traditional fluorescence-based measurements lack the capability to distinguish membrane proteins residing in different organelles. Cutting edge methodologies transcend traditional methods by coupling pH-sensitive fluorophores with total internal reflection fluorescence microscopy (TIRFM). TIRF illumination excites the sample up to approximately 150 nm from the glass-sample interface, thus decreasing background, increasing the signal to noise ratio, and enhancing resolution. The excitation volume in TIRFM encompasses the plasma membrane and nearby organelles such as the peripheral ER. Superecliptic pHluorin (SEP) is a pH sensitive version of GFP. Genetically encoding SEP into the extracellular domain of a membrane protein of interest positions the fluorophore on the luminal side of the ER and in the extracellular region of the cell. SEP is fluorescent when the pH is greater than 6, but remains in an off state at lower pH values. Therefore, receptors tagged with SEP fluoresce when residing in the endoplasmic reticulum (ER) or upon insertion in the plasma membrane (PM) but not when confined to a trafficking vesicle or other organelles such as the Golgi. The extracellular pH can be adjusted to dictate the fluorescence of receptors on the plasma membrane. The difference in fluorescence between TIRF images at neutral and acidic extracellular pH for the same cell corresponds to a relative number of receptors on the plasma membrane. This allows a simultaneous measurement of intracellular and plasma membrane resident receptors. Single vesicle insertion events can also be measured when the extracellular pH is neutral, corresponding to a low pH trafficking vesicle fusing with the plasma membrane and transitioning into a fluorescent state. This versatile technique can be exploited to study localization, expression, and trafficking of membrane proteins.

Labeling the extracellular domain of a membrane protein with a pH sensitive fluorophore, superecliptic pHluorin (SEP), allows subcellular localization, expression, and trafficking to be determined. Imaging SEP-labeled proteins with total internal reflection fluorescence microscopy (TIRFM) enables the quantification of protein levels in the peripheral ER and plasma membrane.

Understanding membrane protein trafficking, assembly, and expression requires an approach that differentiates between those residing in intracellular ...

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how fluorescent recovery after photobleaching (FRAP) and photoactivation techniques can be used to study the spatiotemporal dynamics of proteins in subcellular locations. We also show how these techniques enable straightforward determination of various parameters linked to actin cytoskeletal regulation and cell motility. Moreover, the microinjection of cells is additionally described as an alternative treatment (potentially preceding or complementing the aforementioned photomanipulation techniques) to trigger instantaneous effects of translocated proteins on cell morphology and function. Micromanipulation such as protein injection or local application of plasma membrane-permeable drugs or cytoskeletal inhibitors can serve as powerful tool to record immediate consequences of a given treatment on cell behavior at the single cell and subcellular level. This is exemplified here by immediate induction of lamellipodial cell edge protrusion by the injection of recombinant Rac1 protein, as established a quarter-century ago. In addition, we provide a protocol for determining the turnover of enhanced green fluorescent protein (EGFP)-VASP, an actin filament polymerase prominently accumulating at lamellipodial tips of B16-F1 cells, employing FRAP and including associated data analysis and curve fitting. We also present guidelines for estimating the rates of lamellipodial actin network polymerization, as exemplified by cells expressing EGFP-tagged &#946;-actin. Finally, instructions are given for how to investigate the rates of actin monomer mobility within the cell cytoplasm, followed by actin incorporation at sites of rapid filament assembly, such as the tips of protruding lamellipodia, using photoactivation approaches. None of these protocols is&#160;restricted to components or regulators of the actin cytoskeleton, but can easily be extended to explore in analogous fashion the spatiotemporal dynamics and function of proteins in various different subcellular structures or functional contexts.

We describe how micro- and photomanipulation techniques such as FRAP and photoactivation enable the determination of motility parameters and the spatiotemporal dynamics of proteins within migrating cells. Experimental readouts include subcellular dynamics and turnover of motility regulators or of the underlying actin cytoskeleton.

Fluorescence-based Monitoring of PAD4 Activity via a Pro-fluorescence Substrate Analog

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