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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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 could be similarly leveraged to monitor PAD4 activity. The conversion of arginine to citrulline by PAD4 is expected to greatly reduce its susceptibility to trypsin-mediated hydrolysis. Trypsin has a distinct preference in hydrolyzing the C-terminal amide bonds of both lysine and arginine, with optimal activity near a pH range of 7.5 to 8.5. Thus, all trypsin solutions were prepared at a pH of 7.5 and all assays were conducted at a pH of 8.0. Furthermore, the acetylation of the fluorophore 7-amido-4-methylcoumarin onto the arginine C-terminus serves to &#8220;lock&#8221; its fluorescence in an off-state (Figure 1C). Upon treatment with trypsin, the 7-amido-4-methylcoumarin is uncoupled from the molecule, thus restoring its fluorescence. The change in fluorescence can be readily monitored in solution via its maximum excitation ~440 nm under physiological pH conditions. The enzymatic activity of PAD4 in converting arginine to citrulline should prevent the unmasking of the fluorophore, therefore reducing the fluorescence output. Additionally, the arginine amino terminus was acetylated as a carboxybenzyl moiety to mimic the native peptidic substrate, which lacks the charged amino group. The N-terminus acetylation was crucial since it has been previously reported that neither free arginine nor arginine-methyl ester is citrullinated by PAD4 to any appreciable levels13. This assay, in theory, could be used to monitor the activity of any of the five PAD isozymes. One limitation of the assay may be in cases where different isozymes are found within the same assay solution. All PAD isoforms could potentially convert ZRcoum to its citrulline counterpart. Therefore, selectivity between the isoforms is currently limited. Variations to the assay design in the future will involve the inclusion of amino acids to the N-terminus of arginine. The elongation of the N-terminus may provide the substrate analog with higher selectivity by mimicking the native substrates.
Together, the pro-fluorescent PAD4 substrate analog Z-Arg-7-amido-4-methylcoumarin (ZRcoum) has the proper design features to allow for the monitoring of PAD4 activity. The introduction of the fluorophore to the C-terminal end of arginine within ZRcoum could, in theory, interfere with the ability of the molecule to be a substrate for PAD4. ZRcoum was found to be well accommodated within the binding pocket of PAD4 by analyzing its crystal structure whereby ZRcoum was docked into the known substrate binding pocket (PDB 2DW5)38. The coumarin fluorophore within ZRcoum appears to be well tolerated, an expected finding considering the substrate binding site is housed on a solvent exposed face of the protein (Figure 1B). A possible limitation of the assay as it pertains to the screening of small molecule libraries is that some potential inhibitors may display fluorescent properties that overlap with the fluorescence output from 7-amido-4-methylcoumarin. In this scenario, the fluorescent small molecule would result in a false positive signal. In the future, it is possible that the fluorophore moiety on the substrate analog could be varied to minimize this type of interference.
An additional unique feature of the assay compared to previously developed PAD4 assays is the increase in fluorescence levels due to the presence of a potential PAD4 inhibitor. To have an increase in fluorescence level (as opposed to a decrease) is a beneficial characteristic in terms of screening efforts as it renders the assay less susceptible to false positives. In conclusion, an operationally simple fluorescence-based assay was described for PAD4 activity by designing a pro-fluorescent substrate analog and it was shown that the assay is highly adaptable to conditions typically encountered in high-throughput screening platforms. This assay can be utilized in the future to possibly discover the first truly reversible PAD4 inhibitors.

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) remained in the locked state. Upon addition of excess trypsin/EDTA to the solution, the fluorophores were liberated from ZRcoum but remained mostly unchanged in the presence of PAD4. The addition of EDTA is intended to assist in the cessation of the PAD4 reaction due to its ability to chelate the essential calcium ions. From these data, a 5.2-fold decrease in fluorescence output was observed in the presence of PAD4. Next, the compatibility of the newly developed assay with high-throughput screening platforms was demonstrated. It was of primary interest to analyze four reaction variables that would be most favorable for screening purposes (1) the further miniaturization of the assay volume (2) the possibility of performing the assay at RT (3) the stability of the protein and buffer mixture prior to completing the assay and (4) the potential interference of organic solvents/detergents.
For mid- or high-throughput screening efforts, high-density well plates are commonly utilized in small molecule screens. It was verified that the assay can be performed almost identically in a 384-well plate as compared to a 96-well plate (Figure 2B). In further determining the working conditions for the assay in a high-throughput format, it was important to investigate whether PAD4 activity can be monitored in a reasonable time frame at RT. Since not all liquid-handling instruments have a temperature controller, it can become cumbersome and costly to have an incubation step at elevated temperature. Furthermore, accounting for fluctuation in temperature can be a cause for discrepancy between plates and runs. In order to determine the effect of lower temperatures for the assay described here, a similar assay was performed at RT. It was found that the signals were virtually unchanged between 37 &#176;C and RT (Figure 3). To further optimize the assay performance by minimizing the required incubation time between PAD4 and the substrate, the kinetics of the reaction was analyzed. PAD4 activity was monitored over time both at RT and 37 &#176;C. As expected, it appears that PAD4 is more active at 37 &#176;C (consistent with human physiological temperature) with the reaction mostly complete by 20 min. Satisfyingly, the reaction is not strongly affected by the reduction in temperature down to RT. The reaction proceeds at approximately half the speed as at the elevated temperature and is essentially complete 40-45 min post initiation. From the time course analyses, kinetics constants were calculated and the parameters (KM = 397 &#956;M, kcat = 2.98 sec-1, kcat/kM = 7,400 sec-1 M-1) were found to be similar to other PAD4 substrates.31 With the kinetics of the reaction characterized, all the subsequent assays were reduced to 1 hr from the beginning to the measuring point.
Next, stability of PAD4 was evaluated when dissolved in the reaction buffer over a prolonged period of time. Realistically, high-throughput screening will require reagents to be generally stable for many hours while the instrument is completing the screen. If certain reagents have a relative shelf-life of minutes, it can be quite expensive and time consuming to terminate the screen in order to prepare fresh reagents. Depending on the assay conditions, it is not unusual for some assays to be incompatible with high through-put screening for this very reason. With this idea in mind, the stability of PAD4 in buffer was monitored over several hours. Almost no loss in enzymatic activity was observed throughout the time period, thus indicating that the protein is stable even after sitting at RT for over 15 hr (Figure 4A). Next, it was evaluated whether the assay run under miniaturized conditions and at RT would still appropriately respond to the presence of a PAD4. Upon the incubation of the known PAD4 inhibitor, chloroamidine (Cl-amidine), led to a complete restoration of the fluorescence signal consistent with the full inhibition of PAD4 (Figure 4B). Finally, it was also verified that the presence of the organic solvent DMSO, commonly used in stock solutions of small molecules, up to 3% as final volume, did not interfere with the assay (data not shown). Likewise, the addition of 0.01% Triton X-100 resulted in similar fluorescence trends (data not shown).
Having shown that the assay is tolerant to a variety of conditions that are typically encountered in small molecule drug discovery efforts, a preliminary screening of 80 compounds was conducted and a robust Z&#8217; value of 0.71 was identified. In furthering the potential of the assay, the selectivity of the PAD4/ZRcoum reaction was highlighted by performing the reaction in crude whole cell lysates. E. coli cells containing the GST-PAD4 expression vector were induced with IPTG, similar to the procedure used for the isolation of GST-PAD4. Following the disruption of the cell membrane via sonication, the cellular debris was separated by centrifugation to afford a clarified cell lysate. When the PAD4-containing cell lysate was submitted to the same assay conditions as the purified GST-PAD4 assay, the fluorescence signal varied in agreement with the presence of active PAD4 in solution (Figure 5). Cell lysates contain a number of biomacromolecules at high concentrations that could potentially interfere with the activity of PAD4, the fluorescent properties of ZRcoum, and the trypsin step. The finding that the assay was functional in crude cell lysates indicates that there is a high selectivity in the reaction between PAD4 and ZRcoum. To further demonstrate that the signal decrease in the presence of crude cell lysates was specific to the presence of active PAD4, the effect of Cl-amidine, an established covalent PAD4 inhibitor, was evaluated. The addition of Cl-amidine led to a restoration in the fluorescence signal, consistent with the inactivation of PAD4 (Figure 5). Together, it was demonstrated the assay can be further simplified by utilizing crude cell lysates for the monitoring of PAD4 activity.
<img alt="Figure 1" src="/files/ftp_upload/52114/52114fig1highres.jpg" width="600" /> 
Figure 1. PAD4 and the design of its fluorescence-based assay.&#160;(A) Cartoon representation of the histone assemblies that make up the basic unit of a nucleosome. PAD4 catalyzes the citrullination of a number of arginine residues on histone proteins. (B) ZRcoum overlaid with the native substrate (hidden for clarity) of PAD4 (accession code 2DW5). (C) Representation of the steps in a typical PAD4 assay. Citrullination by PAD4 of the substrate ZRcoum reduces its susceptibility to trypsin-mediated amide hydrolysis, thus causing a change in fluorescence levels. <a href="/files/ftp_upload/52114/52114fig1highres.jpg" target="_blank">Please click here to view a larger version of the figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52114/52114fig2highres.jpg" width="600" /> 
Figure 2. PAD4 assay in 96-well and 384-well plate formats. Fluorescence was measured (excitation 345 nm/emission 465 nm) in both a 96-well format (A) and 384-well format (B) with the designated conditions (37 &#176;C). The addition of protease trypsin leads to a large increase in fluorescence levels in wells without PAD4 and remains virtually unchanged for wells with PAD4. Data are represented as mean + SD (n = 3). <a href="/files/ftp_upload/52114/52114fig2highres.jpg" target="_blank">Please click here to view a larger version of the figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52114/52114fig3highres.jpg" width="600" /> 
Figure 3. Time course analysis of PAD4 assay. Fluorescence was measured (excitation 345 nm/emission 465 nm) periodically in a 96-well plate format over 70 min with the protein being incubated either at 37 &#176;C (A) or at RT (B). Data are represented as mean +/- SD (n = 3). <a href="/files/ftp_upload/52114/52114fig3highres.jpg" target="_blank">Please click here to view a larger version of the figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/52114/52114fig4highres.jpg" width="600" /> 
Figure 4. Protein stability and inhibition. (A) The activity of PAD4 was measured after the designated time intervals. For comparison, the fluorescence levels are shown in the absence of PAD4. Fluorescence levels are shown prior (red) and following (green) the addition of trypsin. (B) Introduction of the known PAD4 inhibitor Cl-amidine (100 &#181;M) led to a restoration of the fluorescence signal in a 384-well plate format at RT. Fluorescence levels are shown prior (red) and following (green) the addition of trypsin. All data are represented as mean +/- SD (n = 3). <a href="/files/ftp_upload/52114/52114fig4highres.jpg" target="_blank">Please click here to view a larger version of the figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/52114/52114fig5highres.jpg" width="300" /> 
Figure 5. Crude cell lysate assay. The assay was performed in the absence and presence of crude cell lysates in a 384-well plate format at RT. Fluorescence levels are shown prior (red) and following (green) the addition of trypsin. Cl-amidine (100 &#181;M) also caused an increase in fluorescence signal under these conditions. All data are represented as mean +/- SD (n = 3). <a href="/files/ftp_upload/52114/52114fig5highres.jpg" target="_blank">Please click here to view a larger version of the figure.</a>

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

Nanyang Technological University

Research

JoVE Journal

Chemistry

9.9K 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

Fluorescence detection methods for microfluidic droplet platforms

The development of microfluidic platforms for performing chemistry and biology has in large part been driven by a range of potential benefits that accompany system miniaturisation. Advantages include the ability to efficiently process nano- to femoto- liter volumes of sample, facile integration of functional components, an intrinsic predisposition towards large-scale multiplexing, enhanced analytical throughput, improved control and reduced instrumental footprints.1
In recent years much interest has focussed on the development of droplet-based (or segmented flow) microfluidic systems and their potential as platforms in high-throughput experimentation.2-4 Here water-in-oil emulsions are made to spontaneously form in microfluidic channels as a result of capillary instabilities between the two immiscible phases. Importantly, microdroplets of precisely defined volumes and compositions can be generated at frequencies of several kHz. Furthermore, by encapsulating reagents of interest within isolated compartments separated by a continuous immiscible phase, both sample cross-talk and dispersion (diffusion- and Taylor-based) can be eliminated, which leads to minimal cross-contamination and the ability to time analytical processes with great accuracy. Additionally, since there is no contact between the contents of the droplets and the channel walls (which are wetted by the continuous phase) absorption and loss of reagents on the channel walls is prevented.
Once droplets of this kind have been generated and processed, it is necessary to extract the required analytical information. In this respect the detection method of choice should be rapid, provide high-sensitivity and low limits of detection, be applicable to a range of molecular species, be non-destructive and be able to be integrated with microfluidic devices in a facile manner. To address this need we have developed a suite of experimental tools and protocols that enable the extraction of large amounts of photophysical information from small-volume environments, and are applicable to the analysis of a wide range of physical, chemical and biological parameters. Herein two examples of these methods are presented and applied to the detection of single cells and the mapping of mixing processes inside picoliter-volume droplets. We report the entire experimental process including microfluidic chip fabrication, the optical setup and the process of droplet generation and detection.

Droplet-based microfluidic platforms are promising candidates for high throughput experimentation since they are able to generate picoliter, self-compartmentalized vessels inexpensively at kHz rates. Through integration with fast, sensitive and high resolution fluorescence spectroscopic methods, the large amounts of information generated within these systems can be efficiently extracted, harnessed and utilized.

The development of microfluidic platforms for performing chemistry and biology has in large part been driven by a range of potential benefits that ...

Bioengineering

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 ...

Biochemistry

cAMP Analysis with cADDis: Single-Cell to High-Throughput Screening

Real-Time cAMP Dynamics in Live Cells Using the Fluorescent cAMP Difference Detector In Situ

cAMP Difference Detector In Situ (cADDis) is a novel biosensor that allows for the continuous measurement of cAMP levels in living cells. The biosensor is created from a circularly permuted fluorescent protein linked to the hinge region of Epac2. This creates a single fluorophore biosensor that displays either increased or decreased fluorescence upon binding of cAMP. The biosensor exists in red and green upward versions, as well as green downward versions, and several red and green versions targeted to subcellular locations. To illustrate the effectiveness of the biosensor, the green downward version, which decreases in fluorescence upon cAMP binding, was used. Two protocols using this sensor are demonstrated: one utilizing a 96-well plate reading spectrophotometer&#160;compatible with high-throughput screening and another utilizing single-cell imaging on a fluorescent microscope. On the plate reader, HEK-293 cells cultured in 96-well plates were stimulated with 10 &#181;M forskolin or 10 nM isoproterenol, which induced rapid and large decreases in fluorescence in the green downward version. The biosensor was used to measure cAMP levels in individual human airway smooth muscle (HASM) cells monitored under a fluorescent microscope. The green downward biosensor displayed similar responses to populations of cells when stimulated with forskolin or isoproterenol. This single-cell assay allows visualization of the biosensor location at 20x and 40x magnification. Thus, this cAMP biosensor is sensitive and flexible, allowing real-time measurement of cAMP in both immortalized and primary cells, and with single cells or populations of cells. These attributes make cADDis a valuable tool for studying cAMP signaling dynamics in living cells.

Author Spotlight: Advancing Real-Time cAMP Detection in Cells Using cADDis Biosensor

The protocol presented in this study illustrates the effectiveness of the cAMP Difference Detector In Situ in measuring cAMP through two methods. One method utilizes a 96-well plate reading spectrophotometer with HEK-293 cells. The other method demonstrates individual HASM cells under a fluorescent microscope.

cAMP Difference Detector In Situ (cADDis) is a novel biosensor that allows for the continuous measurement of cAMP levels in living cells. The biosensor is ...

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

Monitoring Kinase and Phosphatase Activities Through the Cell Cycle by Ratiometric FRET

F&ouml;rster resonance energy transfer (FRET)-based reporters1 allow the assessment of endogenous kinase and phosphatase activities in living cells. Such probes typically consist of variants of CFP and YFP, intervened by a phosphorylatable sequence and a phospho-binding domain. Upon phosphorylation, the probe changes conformation, which results in a change of the distance or orientation between CFP and YFP, leading to a change in FRET efficiency (Fig 1). Several probes have been published during the last decade, monitoring the activity balance of multiple kinases and phosphatases, including reporters of PKA2, PKB3, PKC4, PKD5, ERK6, JNK7, Cdk18, Aurora B9 and Plk19. Given the modular design, additional probes are likely to emerge in the near future10. 
Progression through the cell cycle is affected by stress signaling pathways 11. Notably, the cell cycle is regulated differently during unperturbed growth compared to when cells are recovering from stress12.Time-lapse imaging of cells through the cell cycle therefore requires particular caution. This becomes a problem particularly when employing ratiometric imaging, since two images with a high signal to noise ratio are required to correctly interpret the results. Ratiometric FRET imaging of cell cycle dependent changes in kinase and phosphatase activities has predominately been restricted to sub-sections of the cell cycle8,9,13,14.
Here, we discuss a method to monitor FRET-based probes using ratiometric imaging throughout the human cell cycle. The method relies on equipment that is available to many researchers in life sciences and does not require expert knowledge of microscopy or image processing.

FRET-based reporters are increasingly used to monitor kinase and phosphatase activities in live cells. Here we describe a method on how to use FRET-based reporters to assess cell cycle-dependent changes in target phosphorylation.

F&ouml;rster resonance energy transfer (FRET)-based reporters1 allow the assessment of endogenous kinase and phosphatase activities in living cells. Such ...

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 ...

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

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 ...

Real-Time Monitoring of Aurora kinase A Activation using Conformational FRET Biosensors in Live Cells

Epithelial cancers are often hallmarked by the overexpression of the Ser/Thr kinase Aurora A/AURKA. AURKA is a multifunctional protein that activates upon its autophosphorylation on Thr288. AURKA abundance peaks in mitosis, where it controls the stability and the fidelity of the mitotic spindle, and the overall efficiency of mitosis. Although well characterized at the structural level, a consistent monitoring of the activation of AURKA throughout the cell cycle is lacking. A possible solution consists in using genetically-encoded F&#246;rster&#8217;s Resonance Energy Transfer (FRET) biosensors to gain insight into the autophosphorylation of AURKA with sufficient spatiotemporal resolution. Here, we describe a protocol to engineer FRET biosensors detecting Thr288 autophosphorylation, and how to follow this modification during mitosis. First, we provide an overview of possible donor/acceptor FRET pairs, and we show possible cloning and insertion methods of AURKA FRET biosensors in mammalian cells. Then, we provide a step-by-step analysis for rapid FRET measurements by fluorescence lifetime imaging microscopy (FLIM) on a custom-built setup. However, this protocol is also applicable to alternative commercial solutions available. We conclude by considering the most appropriate FRET controls for an AURKA-based biosensor, and by highlighting potential future improvements to further increase the sensitivity of this tool.

The activation of the multifunctional Ser/Thr kinase AURKA is hallmarked by its autophosphorylation on Thr288. Fluorescent probes relying on FRET can discriminate between its inactive and activated states. Here, we illustrate some strategies for probe engineering, together with a rapid FRET protocol to follow the kinase activation throughout mitosis.

Epithelial cancers are often hallmarked by the overexpression of the Ser/Thr kinase Aurora A/AURKA. AURKA is a multifunctional protein that activates upon ...

Time-Resolved Forster Resonance Energy Transfer for Monitoring Protein Phosphorylation in Cells

Source: Padros, J. et al., <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/62915">Time-resolved F&#246;rster Resonance Energy Transfer Assays for Measurement of Endogenous Phosphorylated STAT Proteins in Human Cells.</a> J. Vis. Exp. (2021).
This video describes the time-resolved F&#246;rster resonance energy transfer or TR-FRET assay to determine endogenous protein phosphorylation with the help of donor and acceptor fluorophores tagged on antibodies specific to a test protein. This technique is designed to measure the phosphorylation of test proteins in cell lysates.

Source: Padros, J. et al., Time-resolved F&#246;rster Resonance Energy Transfer Assays for Measurement of Endogenous Phosphorylated STAT Proteins in Human ...

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

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