The protocol was adopted from a previous publication11. T. Z., D. Ch., and J. &#268;. were supported by the Ministry of Education, Youth and Sports of the Czech Republic within the National Sustainability Program I (NPU I), grant number LO1415. J. &#268;. was also supported by GA CR, Grant number 18-24397S. Access to instruments and other facilities was supported by the Czech research infrastructure for systems biology C4SYS (project no LM2015055). M. A. S. was supported by a grant from the Russian Science Foundation [no. 14-14-00904].

1. Cyanobacteria Cultivation

<ol>
	<li>Cultivate Synechocystis cells in Erlenmeyer flasks or in photobioreactors10,19 in buffered BG11 medium20 to maintain a pH of &#60; 10 (e.g., using 17 mM HEPES10). 
	NOTE: Standard cultivation conditions require a controlled temperature (typically, 30 &#176;C, the optimal temperature is 35 &#176;C)21, illumination (typically, a white light of an intensity up to 800 &#181;mol[photons]/[m2&#183;s])21, and a CO2 supply (in the 400-mL flat-panel photobioreactor, the growth saturating CO2 concentration is 1,700 ppm)21.</li>
	<li>To determine the culture density, measure the culture optical density spectrophotometrically at 730 nm (OD730) using a cuvette with a light path of 1 cm. Alternatively, count the cells under a microscope using a hemocytometer or an automated cell counter.</li>
</ol>
2. Samples Preparation
<ol>
	<li>Work under low irradiance to prevent phycobiliprotein degradation.</li>
	<li>Harvest 3 x 1 mL of culture suspension to safe-lock tubes. Use triplicates for the estimation of technical errors in the measurement. To perform the culture sampling under sterile conditions, harvest the cells in a laminar hood and follow the proper working and safety practices.</li>
	<li>Centrifuge the cells at 15,000 x g at laboratory temperature for 5 min. Pay attention to the properly balanced centrifuge rotor. After centrifugation, discard the supernatants. Be sure not to disturb the pellet.</li>
	<li>Put the samples in a freezer. For long-term storage, keep the samples at -80 &#176;C. This is necessary to prevent phycobiliprotein degradation. For short-term storage, -20 &#176;C is sufficient.</li>
	<li>Freeze-dry the samples overnight. For a proper freeze-drying process, keep the temperature of the freeze-dryer condenser below -60 &#176;C and the pressure in the freeze-dryer chamber around 1 hPa.</li>
	<li>After finishing the freeze-drying cycle, close the tube as soon as possible to prevent the reabsorption of water from the air.</li>
</ol>
3. Cell Homogenization and Pigments Extraction
<ol>
	<li>Add four pieces of glass beads (with a diameter of 2 mm) to each sample tube and close the tubes. 
	NOTE: When the lid of the safe-lock tube used for homogenization is too thin, it can break during the homogenization; therefore, only safe-lock tubes with strong lids are recommended for this protocol.</li>
	<li>Homogenize the samples with the glass beads for 15 s on a homogenizer at laboratory temperature. 
	NOTE: A properly homogenized sample is spread over the whole inner surface of the safe-lock tubes.</li>
	<li>Add 1 mL of PBS buffer (pH 7.4), precooled to 4 &#176;C, to the samples in order to extract phycobiliproteins. 
	NOTE: From this step on, keep the samples on ice to prevent the degradation of the extracted proteins. This is critical.</li>
	<li>Mix the samples with PBS for 5 s on the homogenizer at laboratory temperature. 
	NOTE: After mixing, the samples are greenish.</li>
	<li>After mixing, keep the samples on ice for 60 min. Cover the ice bath with a lid to prevent pigments degradation.</li>
	<li>After 60 min of phycobiliprotein extraction, centrifuge the samples at 15,000 x g at 4 &#176;C for 5 min. Pay attention to properly balance the centrifuge rotor. 
	NOTE: After the centrifugation, the supernatant has a cyan blue color.</li>
</ol>
4. Phycobiliprotein Quantification
<ol>
	<li>Before the spectrophotometric measurement, calibrate the spectrophotometer to the baseline using the PBS buffer as a blank.</li>
	<li>Once the spectrophotometer is calibrated, discard the PBS from one spectrophotometer cuvette and pipette the supernatant with extracted phycobiliproteins instead of with the discarded buffer.</li>
	<li>Quantify the phycobiliprotein concentration spectrophotometrically, using a slit width of 0.5 nm.
	<ol>
		<li>Measure the absorbance of the phycocyanobilins in phycocyanin and allophycocyanin against the PBS buffer blank at 615 nm (A615) and 652 nm (A652), respectively, and measure the absorbance of the cellular debris at 720 nm (A720).</li>
		<li>Calculate the concentration of phycocyanin and allophycocyanin according to the equations (1) and (2) of Bennett and Bogorad12: 
		<img alt="Equation 1" src="/files/ftp_upload/58076/58076eq1.jpg" /> 
		<img alt="Equation 2" src="/files/ftp_upload/58076/58076eq2.jpg" /> 
		NOTE: A615, A652, and A720 should fit to the linear absorbance range of the spectrophotometer. If necessary, dilute the sample with PBS buffer.</li>
	</ol>
	</li>
	<li>Recalculate the pigment concentration in the original samples. The pigment concentration in the sample corresponds directly with the results of equations (1) and (2) when 1 mL of culture and 1 mL of the extraction buffer are used for the analysis. In case of using different volumes of cyanobacteria samples and/or PBS buffer, the final pigment concentration should be calculated according to equation (3): 
	<img alt="Equation 3" src="/files/ftp_upload/58076/58076eq3.jpg" />&#160; &#160;(3) 
	NOTE: In case of a normalizing phycobiliprotein content per cell dry weight, the final pigment concentration should be calculated according to equation (4)<img alt="Equation 4" src="/files/ftp_upload/58076/58076eq4.jpg" />&#160; (4)</li>
</ol>
5. Determination of the Cell Dry Weight (Optional)
<ol>
	<li>Weigh three empty safe-lock tubes on analytical balances. 
	CAUTION: It is critical that the safe-lock tubes are dry. In case of storing the tubes in a wet environment, dry the tubes for several hours in a freeze dryer before weighing them. Manipulate the tubes gently and only with powder-free gloved hands to avoid any contact between the material and the researcher&#39;s fingers. Keep all holders and centrifuge clean to avoid the transfer of any material to the tubes.</li>
	<li>Sample 1 x 15 mL of culture suspension to 15-mL conical tube.</li>
	<li>Centrifuge the culture suspension in the 15-mL conical tubes at 4,000 x g at laboratory temperature for 10 min. Balance the centrifuge rotor with three additional 15-mL conical tubes, each containing 15 mL of water. After centrifugation, discard 12 mL of the supernatant.</li>
	<li>Resuspend the pellet in the remaining supernatant and transfer 3 x 1 mL of the mixture to three 1.5-mL safe-lock tubes with a pipette. In case some leftovers of the pellet remain in the 15-mL conical tube, add 1.5 mL of deionized water to the conical tube, vortex or shake the tube to resuspend the remaining pellet, and transfer 3 x 0.5 mL of the mixture to the three 1.5-mL safe-lock tubes (already containing 3 x 1 mL of the pellet) with a pipette. 
	NOTE: Dividing the pellet over three 1.5-mL safe-lock tubes will allow the estimation of any technical error of the dry weight measurement.</li>
	<li>Centrifuge the cells in 1.5-mL safe-lock tubes at 15,000 x g at laboratory temperature for 5 min. Balance the centrifuge rotor with an additional 1.5-mL safe-lock tube that contains 1 mL of water. After the centrifugation, discard the supernatants.</li>
	<li>Put the samples in the freezer. For long-term storage, keep the samples at -80 &#176;C; for short-term storing, -20 &#176;C is sufficient.</li>
	<li>Freeze-dry the samples overnight. For a proper freeze-drying process, keep the temperature of the freeze dryer condenser below -90 &#176;C and the pressure in the freeze-dryer chamber around 1 hPa.</li>
	<li>After freeze-drying, close the tubes and weigh the samples on analytical balances. 
	NOTE: The dry weight value can be used for the normalization of the phycobiliprotein content per cell dry weight.</li>
</ol>

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The authors have nothing to disclose.

This protocol describes a simple, fast, and reproducible method for the quantification of phycobiliprotein content in the model cyanobacterium Synechocystis. Several methods of cell homogenization, protein extraction, and phycocyanin and allophycocyanin quantification are compared, and the final protocol represents a combination of the optimal steps of every single procedure. As representative data, the content of phycobiliproteins was quantified in Synechocystis cells under increasing light intensity. ...

fPhycobiliproteins are water-soluble pigment-protein complexes that represent major components of the light-harvesting antennae in prokaryotic cyanobacteria (Cyanophyta) and several eukaryotic taxa (Glaucophyta, Rhodophyta, and Cryptophyta)1. They occur mainly as supramolecular complexes called phycobilisomes and they are typically attached to the surface of the photosynthetic membranes on the stromal side, with the exception of Cryptophyta, where the phycobiliproteins are localized in the thylakoid lumen2. Four types of phycobiliproteins have been identified up to date: the core allophycocyanin, and the peripheral phycocyanin, phycoerythrin, and phycoerythrocyanin1. As the main light-harvesting complexes, phycobilisomes represent one of the crucial factors of algae and cyanobacteria mass cultures productivity. It has been demonstrated that phycobilisomes truncation can enhance biomass accumulation under strong light3. On the other hand, under modest or low irradiance, the antenna truncation resulted in growth rates and biomass accumulation reduction3,4. Phycobiliproteins are commercially used as food colorants, pharmaceuticals, and food additives, in the cosmetic industry, and as fluorescence probes with applications in flow cytometry, fluorescent immunoassays, and fluorescence microscopy5.
This protocol focuses on the quantitative determination of phycobiliproteins in the model cyanobacterium Synechocystis. Cyanobacteria are the earliest oxygenic photosynthetic autotrophs; they have been forming the Earth&#39;s biosphere for more than 2.4 billion years6. They play a crucial role in global biogeochemical cycles of nitrogen, carbon, oxygen, and other elements. Among cyanobacteria, a unicellular strain Synechocystis gained a unique position since it was the first cyanobacterium with the entire genome sequenced7,8, it is naturally transformable by exogenous DNA9, and it performs stable and relatively fast growth10,11. In Synechocystis, the core antenna component, allophycocyanin, is associated with the integral membrane proteins, and the attached phycocyanin is located on the thylakoid membrane periphery.
Several methods for phycobiliprotein extraction and quantification are compared within this protocol. The final extraction procedure was successfully applied to Synechocystis, as well as to other cyanobacteria strains, including Cyanothece sp., Synechococcuselongatus, Spirulina sp., Arthrospira sp., and Nostoc sp., and it was also successfully applied to red algae Porphyridium cruentum. Therefore, the method developed in this protocol can be considered as a universal method for phycobiliprotein extraction. Even though some of the tested extraction methods resulted in higher total protein yields, the here described extraction procedure provided the highest phycobiliprotein yields together with the lowest content of chlorophyll a residue in the phycobiliprotein extract. Reducing the content of chlorophyll a was essential for the correct phycocyanin and allophycocyanin spectrophotometric quantification.
The phycobiliprotein absorption spectra can vary significantly among various algae and cyanobacteria species12,13,14,15,16,17 and even among several strains of a single cyanobacteria genus18. Therefore, the specific wavelengths and absorption coefficients as used for the determination of phycocyanin and allophycocyanin in Synechocystis are not generally applicable to other strains. Additionally, Synechocystis does not contain phycoerythrin and phycoerythrocyanin that can be found in some other algae and cyanobacteria. For the purpose of the determination of phycobiliproteins in strains other than Synechocystis, it is recommended to evaluate the spectrophotometric equations for each strain individually.
Although the protocol contains two longer steps (overnight freeze-drying of the cellular pellets and 1-hour protein extraction), the total labor time for the phycobiliproteins quantification is no longer than 2 hours.

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			<td height="21" style="height:21px;width:407px;">Synechocystis sp. PCC 6803</td>
			<td style="width:417px;">Institut Pasteur, Paris, France</td>
			<td style="width:137px;">6803</td>
			<td style="width:627px;">Cyanobacterium strain</td>
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			<td height="21" style="height:21px;">Roti-CELL PBS</td>
			<td>Carl Roth GmbH + Co. KG, Karlsruhe, Germany</td>
			<td>9143.1</td>
			<td>Phosphate-Buffered Saline (PBS) solution, pH 7.4</td>
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			<td height="21" style="height:21px;">Eppendorf safe-lock tubes&#160;</td>
			<td style="width:417px;">Eppendorf, Hamburk, Germany</td>
			<td>30120086</td>
			<td>Safe-lock tubes 1.5 ml</td>
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			<td height="21" style="height:21px;">VWR 80-Place Storage System</td>
			<td style="width:417px;">VWR International, Radnor, Pennsylvania, USA</td>
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			<td>Holder for safe-lock tubes&#160;</td>
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			<td>Mettler-Toledo, Columbus, Ohio, USA</td>
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			<td>Mettler-Toledo, Columbus, Ohio, USA</td>
			<td>30389272</td>
			<td>Pipette tips</td>
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			<td height="21" style="height:21px;">Rotina 420R</td>
			<td>Hettich, Kirchlengern, Germany</td>
			<td>4701</td>
			<td>Refrigerated centrifuge for 1.5 ml safe-lock tubes and 15 ml conical centrifuge tubes</td>
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			<td height="21" style="height:21px;">LCexv 4010</td>
			<td>Liebherr, Bulle, Switzerland</td>
			<td>9005382197172</td>
			<td>Refrigerator and freezer -20&#160;&#176;C</td>
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			<td height="21" style="height:21px;width:407px;">Revco ExF -86&#176;C Upright Ultra-Low Temperature Freezer</td>
			<td style="width:417px;">Thermo Fisher Scientific, Waltham, Massachusetts, USA</td>
			<td style="width:137px;">EXF24086V&#160;</td>
			<td>Freezer -80&#160;&#176;C</td>
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			<td>LaboGene, Liller&#248;d, Denmark</td>
			<td>7.001.000.615</td>
			<td>Freeze dryer&#160;</td>
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			<td height="21" style="height:21px;">UV-2600</td>
			<td>Shimadzu, Kyoto, Japan</td>
			<td>UV-2600</td>
			<td>Spectrophotometer&#160;</td>
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			<td height="21" style="height:21px;">Hellma absorption cuvettes, semi Micro</td>
			<td>Sigma-Aldrich, St. Louis, Missouri, USA</td>
			<td>Z600288&#160;</td>
			<td>VIS/UV-VIS semi-micro cuvettes 0.75-1.5 ml, spectral range 200-2500 nm&#160;</td>
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			<td height="21" style="height:21px;">Silamat S6</td>
			<td>Ivoclar Vivadent, Schaan, Liechtenstein</td>
			<td>602286WU</td>
			<td>Homogenizer&#160;</td>
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			<td height="21" style="height:21px;">Solid-glass beads</td>
			<td>Sigma-Aldrich, St. Louis, Missouri, USA</td>
			<td>Z273627</td>
			<td>Glass bead of the diameter 2&#160;mm</td>
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			<td height="21" style="height:21px;">CPA225D-0CE</td>
			<td>Sartorius AG, G&#246;ttingen, Germany</td>
			<td>SECURA225D-1OBR</td>
			<td>Analytical balances</td>
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		<tr height="21">
			<td height="21" style="height:21px;">C-Phycocyanin from Spirulina sp.&#160;</td>
			<td>Sigma-Aldrich, St. Louis, Missouri, USA</td>
			<td>P2172</td>
			<td>Phycocyanin standard</td>
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		<tr height="21">
			<td height="21" style="height:21px;">Allophycocyanin</td>
			<td>Sigma-Aldrich, St. Louis, Missouri, USA</td>
			<td>A7472</td>
			<td>Allophycocyanin standard</td>
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		<tr height="21">
			<td height="21" style="height:21px;width:407px;">Bicinchoninic Acid Kit&#160;</td>
			<td style="width:417px;">Sigma-Aldrich, St. Louis, Missouri, USA</td>
			<td style="width:137px;">BCA1, B9643</td>
			<td>Complete kit for total proteins determination</td>
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		<tr height="21">
			<td height="21" style="height:21px;width:407px;">AlgaeTron&#160;</td>
			<td style="width:417px;">Photon System Instruments Ltd., Dr&#225;sov, Czech Republic</td>
			<td style="width:137px;">AG 130-ECO&#160;</td>
			<td>Cultivation chamber for E. flasks, with controllable light and atmosphere</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:407px;">Photobioreactor</td>
			<td style="width:417px;">Photon System Instruments Ltd., Dr&#225;sov, Czech Republic</td>
			<td style="width:137px;">FMT-150</td>
			<td>Cultivation equipment for cyanobacteria and algae with completely controllable environment</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Cellometer&#160;</td>
			<td style="width:417px;">Nexcelom Bioscience, Lawrence, Massachusetts, USA</td>
			<td style="width:137px;">Auto M10</td>
			<td>Cell counter</td>
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		<tr height="21">
			<td height="21" style="height:21px;width:407px;">Corning 15 mL centrifuge tubes</td>
			<td style="width:417px;">Sigma-Aldrich, St. Louis, Missouri, USA</td>
			<td style="width:137px;">CLS430791&#160;</td>
			<td>15 ml Centrifuge tube for dry weigth sampling</td>
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		<tr height="21">
			<td height="21" style="height:21px;width:407px;">Herasafe KS</td>
			<td style="width:417px;">Thermo Fisher Scientific, Waltham, Massachusetts, USA</td>
			<td style="width:137px;">51024579</td>
			<td>Laminar flow hood</td>
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spectrophotometric determination of phycobiliprotein content in cyanobacterium synechocystis

This is a simple protocol for the quantitative determination of phycobiliprotein content in the model cyanobacterium Synechocystis. Phycobiliproteins are the most important components of phycobilisomes, the major light-harvesting antennae in cyanobacteria and several algae taxa. The phycobilisomes of Synechocystis contain two phycobiliproteins: phycocyanin and allophycocyanin. This protocol describes a simple, efficient, and reliable method for the quantitative determination of both phycocyanin and allophycocyanin in this model cyanobacterium. We compared several methods of phycobiliprotein extraction and spectrophotometric quantification. The extraction procedure as described in this protocol was also successfully applied to other cyanobacteria strains such as Cyanothece sp., Synechococcuselongatus, Spirulina sp., Arthrospira sp., and Nostoc sp., as well as to red algae Porphyridium cruentum. However, the extinction coefficients of specific phycobiliproteins from various taxa can differ and it is, therefore, recommended to validate the spectrophotometric quantification method for every single strain individually. The protocol requires little time and can be performed in any standard life science laboratory since it requires only standard equipment.

For the initial method tests, Synechocystis was cultivated as batch cultures in Erlenmeyer flasks on a shaker in BG11 cultivation medium20 (supplemented with 17 mM HEPES) at 25 &#176;C, under a warm white light of an intensity of 50 &#181;mol(photons)/(m2&#183;s) and with 1% CO2 in the culturing atmosphere. During the cultivation, the cultures were sampled to safe-lock tubes and centrifuged (15,000 x g at laboratory temperatu...

Watch this Scientific Journal Video about Spectrophotometric Determination of Phycobiliprotein Content in Cyanobacterium Synechocystis at JoVE.com

Spectrophotometric Determination of Phycobiliprotein Content in Cyanobacterium Synechocystis

Here, we present a protocol to quantitatively determine phycobiliprotein content in the cyanobacterium Synechocystis using a spectrophotometric method. The extraction procedure was also successfully applied to other cyanobacteria and algae strains; however, due to variations in pigment absorption spectra, it is necessary to test the spectrophotometric equations for each strain individually.

Spectrophotometric Determination of Phycobiliprotein Content in Cyanobacterium Synechocystis

This is a simple protocol for the quantitative determination of phycobiliprotein content in the model cyanobacterium Synechocystis. Phycobiliproteins are ...

spectrophotometric-determination-phycobiliprotein-content

Research

JoVE Journal

Biochemistry

15.1K Views.  Czech Academy of Sciences. Here, we present a protocol to quantitatively determine phycobiliprotein content in the cyanobacterium Synechocystis using a spectrophotometric method. The extraction procedure was also successfully applied to other cyanobacteria and algae strains; however, due to variations in pigment absorption spectra, it is necessary to test the spectrophotometric equations for each strain individually.

Video: Spectrophotometric Determination of Phycobiliprotein Content in Cyanobacterium Synechocystis

Automated Acoustic Dispensing for the Serial Dilution of Peptide Agonists in Potency Determination Assays

As with small molecule drug discovery, screening for peptide agonists requires the serial dilution of peptides to produce concentration-response curves. Screening peptides affords an additional layer of complexity as conventional tip-based sample handling methods expose peptides to a large surface area of plasticware, providing an increased opportunity for peptide loss via adsorption. Preventing excessive exposure to plasticware reduces peptide loss via adherence to plastics and thus minimizes inaccuracies in potency prediction, and we have previously described the benefits of non-contact acoustic dispensing for in vitro high-throughput screening of peptide agonists1. Here we discuss a fully integrated automation solution for non-contact acoustic preparation of peptide serial dilutions in microtiter plates utilizing the example of screening for peptide agonists at the mouse glucagon-like peptide-1 receptor (GLP-1R). Our methods allow for high-throughput cell-based assays to screen for agonists and are easily scalable to support increased sample throughput, or to allow for increased numbers of assay plate copies (e.g., for a panel of more target cell lines).

Peptide adsorption to plasticware during traditional tip-based serial dilutions can significantly impact potency determination and confound the understanding of structure-activity relationships used for lead identification and lead optimization phases of drug discovery. Here methods for automated acoustic non-contact serial dilution of peptide samples are described.

As with small molecule drug discovery, screening for peptide agonists requires the serial dilution of peptides to produce concentration-response curves. ...

Determination of the Glycogen Content in Cyanobacteria

Cyanobacteria accumulate glycogen as a major intracellular carbon and energy storage&#160;during photosynthesis. Recent developments in research have highlighted complex mechanisms of glycogen metabolism, including the diel cycle of biosynthesis and catabolism, redox regulation, and the involvement of non-coding RNA. At the same time, efforts are being made to redirect carbon from glycogen to desirable products in genetically engineered cyanobacteria to enhance product yields. Several methods are used to determine the glycogen contents in cyanobacteria, with variable accuracies and technical complexities. Here, we provide a detailed protocol for the reliable determination of the glycogen content in cyanobacteria that can be performed in a standard life science laboratory. The protocol entails the selective precipitation of glycogen from the cell lysate and the enzymatic depolymerization of glycogen to generate glucose monomers, which are detected by a glucose oxidase-peroxidase (GOD-POD) enzyme coupled assay. The method has been applied to Synechocystis sp. PCC 6803 and Synechococcus sp. PCC 7002, two model cyanobacterial species that are widely used in metabolic engineering. Moreover, the method successfully showed differences in the glycogen contents between the wildtype and mutants defective in regulatory elements or glycogen biosynthetic genes.

Here, we present a reliable and easy assay to measure the glycogen content in cyanobacterial cells. The procedure entails precipitation, selectable depolymerization, and the detection of glucose residues. This method is suitable for both wildtype and genetically engineered strains and can facilitate the metabolic engineering of cyanobacteria.

Cyanobacteria accumulate glycogen as a major intracellular carbon and energy storage&#160;during photosynthesis. Recent developments in research have ...

High Throughput, Absolute Determination of the Content of a Selected Protein at Tissue Levels Using Quantitative Dot Blot Analysis (QDB)

Lacking a convenient, quantitative, high throughput immunoblot method for absolute determination of the content of a specific protein at cellular and tissue level significantly hampers the progress in proteomic research. Results derived from currently available immunoblot techniques are also relative, preventing any efforts to combine independent studies with a large-scale analysis of protein samples. In this study, we demonstrate the process of quantitative dot blot analysis (QDB) to achieve absolute quantification in a high throughput format. Using a commercially available protein standard, we are able to determine the absolute content of capping actin protein, gelsolin-like (CAPG) in protein samples prepared from three different mouse tissues (kidney, spleen, and prostate) together with a detailed explanation of the experimental details. We propose the QDB analysis as a convenient, quantitative, high throughput immunoblot method of absolute quantification of individual proteins at the cellular and tissue level. This method will substantially aid biomarker validation and pathway verification in various areas of biological and biomedical research.

High Throughput, Absolute Determination of the Content of a Selected Protein at Tissue Levels Using Quantitative Dot Blot Analysis QDB

Here we demonstrate a detailed process of quantitative dot blot analysis (QDB) by determining the absolute content of a targeted protein, capping actin protein, gelsolin-like (CAPG), in three different mouse tissues. We demonstrate a high throughput, convenient, quantitative immunoblot technique for biomarker validation at the cellular and tissue level.

Lacking a convenient, quantitative, high throughput immunoblot method for absolute determination of the content of a specific protein at cellular and ...

A Colorimetric Method for Measuring Iron Content in Plants

Iron, one of the most important micronutrients in living organisms, is involved in basic processes, such as respiration and photosynthesis. Iron content is rather low in all organisms, amounting in plants to about 0.009% of dry weight. To date, one of the most accurate methods for measuring iron concentration in plant tissues is flame absorption atomic spectroscopy. However, this approach is time-consuming and expensive and requires specific equipment not commonly found in plant laboratories. Therefore, a simpler, yet accurate method that can be routinely used is needed. The colorimetric Prussian Blue method is regularly used for qualitative iron staining in animal and plant histological sections. In this study, we adapted the Prussian Blue method for quantitative measurements of iron in tobacco leaves. We validated the accuracy of this method using both atomic spectroscopy and Prussian Blue staining to measure iron content in the same samples and found a linear regression (R2 = 0.988) between the two procedures. We conclude that the Prussian Blue method for quantitative iron measurement in plant tissues is precise, simple, and inexpensive. However, the linear regression presented here may not be appropriate for other plant species, due to potential interactions between the sample and the reagent. Establishment of a regression curve is thus needed for different plant species.

We present a simple and reliable protocol for measuring iron content in plant tissues using the colorimetric Prussian Blue method.

Iron, one of the most important micronutrients in living organisms, is involved in basic processes, such as respiration and photosynthesis. Iron content ...

Using High Content Imaging to Quantify Target Engagement in Adherent Cells

Quantitating the interaction of small molecules with their intended protein target is critical for drug development, target validation and chemical probe validation. Methods that measure this phenomenon without modification of the protein target or small molecule are particularly valuable though technically challenging. The cellular thermal shift assay (CETSA) is one technique to monitor target engagement in living cells. Here, we describe an adaptation of the original CETSA protocol, which allows for high throughput measurements while retaining subcellular localization at the single cell level. We believe this protocol offers important advances to the application of CETSA for in-depth characterization of compound-target interaction, especially in heterogeneous populations of cells.

Measurements of drug target engagement are central to effective drug development and chemical probe validation. Here, we detail a protocol for measuring drug-target engagement using high content imaging in a microplate-compatible adaption of the cellular thermal shift assay (CETSA).

Quantitating the interaction of small molecules with their intended protein target is critical for drug development, target validation and chemical probe ...

Enrichment of Native Lipoprotein Particles with microRNA and Subsequent Determination of Their Absolute/Relative microRNA Content and Their Cellular Transfer Rate

Lipoprotein particles are predominately transporters of lipids and cholesterol in the bloodstream. Furthermore, they contain small amounts of strands of noncoding microRNA (miRNA). In general, miRNA alters the protein expression profile due to interactions with messenger-RNA (mRNA). Thus, knowledge of the relative and absolute miRNA content of lipoprotein particles is essential to estimate the biological effect of cellular particle uptake. Here, a quantitative real-time polymerase chain reaction (qPCR)-based protocol is presented to determine the absolute miRNA content of lipoprotein particles&#8212;exemplified shown for native and miRNA-enriched lipoprotein particles. The relative miRNA content is quantified using multiwell microfluidic array cards. Furthermore, this protocol allows scientists to estimate the cellular miRNA and, thus, the lipoprotein particle uptake rate. A significant increase of the cellular miRNA level is observable when using high-density lipoprotein (HDL) particles artificially loaded with miRNA, whereas incubation with native HDL particles yields no significant effect due to their rather low miRNA content. In contrast, the cellular uptake of low-density lipoprotein (LDL) particles&#8212;neither with native miRNA nor artificially loaded with it&#8212;did not alter the cellular miRNA level.

Here, a quantitative real-time polymerase chain reaction-based protocol is presented for the determination of the native micro-RNA content (absolute/relative) of lipoprotein particles. In addition, a method for increasing the micro-RNA level, as well as a method for determining the cellular uptake rate of lipoprotein particles, is demonstrated.

Lipoprotein particles are predominately transporters of lipids and cholesterol in the bloodstream. Furthermore, they contain small amounts of strands of ...

Spectrophotometric Screening for Potential Inhibitors of Cytosolic Glutathione S-Transferases

Glutathione S-transferases (GSTs) are metabolic enzymes responsible for the elimination of endogenous or exogenous electrophilic compounds by glutathione (GSH) conjugation. In addition, GSTs are regulators of mitogen-activated protein kinases (MAPKs) involved in apoptotic pathways. Overexpression of GSTs is correlated with decreased therapeutic efficacy among patients undergoing chemotherapy with electrophilic alkylating agents. Using GST inhibitors may be a potential solution to reverse this tendency and augment treatment potency. Achieving this goal requires the discovery of such compounds, with an accurate, quick, and easy enzyme assay. A spectrophotometric protocol using 1-chloro-2,4-dinitrobenzene (CDNB) as the substrate is the most employed method in the literature. However, already described GST inhibition experiments do not provide a protocol detailing each stage of an optimal inhibition assay, such as the measurement of the Michaelis-Menten constant (Km) for CDNB or indication of the employed enzyme concentration, crucial parameters to assess the inhibition potency of a tested compound. Hence, with this protocol, we describe each step of an optimized spectrophotometric GST enzyme assay, to screen libraries of potential inhibitors. We explain the calculation of both the half-maximal inhibitory concentration (IC50) and the constant of inhibition (Ki)&#8212;two characteristics used to measure the potency of an enzyme inhibitor. The method described can be implemented using a pool of GSTs extracted from cells or pure recombinant human GSTs, namely GST alpha 1 (GSTA1), GST mu 1 (GSTM1) or GST pi 1 (GSTP1). However, this protocol cannot be applied to GST theta 1 (GSTT1), as CDNB is not a substrate for this isoform. This method was used to test the inhibition potency of curcumin using GSTs from equine liver. Curcumin is a molecule exhibiting anti-cancer properties and showed affinity towards GST isoforms after in silico docking predictions. We demonstrated that curcumin is a potent competitive GST inhibitor, with an IC50 of 31.6 &#177; 3.6 &#181;M and a Ki of 23.2 &#177; 3.2 &#181;M. Curcumin has potential to be combined with electrophilic chemotherapy medication to improve its efficacy.

Glutathione S-transferases (GSTs) are detoxification enzymes involved in the metabolism of numerous chemotherapeutic drugs. Overexpression of GSTs is correlated with cancer chemotherapy resistance. One way to counter this phenotype is to use inhibitors. This protocol describes a method using a spectrophotometric assay to screen for potential GST inhibitors.

Glutathione S-transferases (GSTs) are metabolic enzymes responsible for the elimination of endogenous or exogenous electrophilic compounds by glutathione ...

Enabling Real-Time Compensation in Fast Photochemical Oxidations of Proteins for the Determination of Protein Topography Changes

Fast photochemical oxidation of proteins (FPOP) is a mass spectrometry-based structural biology technique that probes the solvent-accessible surface area of proteins. This technique relies on the reaction of amino acid side chains with hydroxyl radicals freely diffusing in solution. FPOP generates these radicals in situ by laser photolysis of hydrogen peroxide, creating a burst of hydroxyl radicals that is depleted on the order of a microsecond. When these hydroxyl radicals react with a solvent-accessible amino acid side chain, the reaction products exhibit a mass shift that can be measured and quantified by mass spectrometry. Since the rate of reaction of an amino acid depends in part on the average solvent accessible surface of that amino acid, measured changes in the amount of oxidation of a given region of a protein can be directly correlated to changes in the solvent accessibility of that region between different conformations (e.g., ligand-bound versus ligand-free, monomer vs. aggregate, etc.) FPOP has been applied in a number of problems in biology, including protein-protein interactions, protein conformational changes, and protein-ligand binding. As the available concentration of hydroxyl radicals varies based on many experimental conditions in the FPOP experiment, it is important to monitor the effective radical dose to which the protein analyte is exposed. This monitoring is efficiently achieved by incorporating an inline dosimeter to measure the signal from the FPOP reaction, with laser fluence adjusted in real-time to achieve the desired amount of oxidation. With this compensation, changes in protein topography reflecting conformational changes, ligand-binding surfaces, and/or protein-protein interaction interfaces can be determined in heterogeneous samples using relatively low sample amounts.

Fast photochemical oxidation of proteins is an emerging technique for the structural characterization of proteins. Different solvent additives and ligands have varied hydroxyl radical scavenging properties. To compare the protein structure in different conditions, real-time compensation of hydroxyl radicals generated in the reaction is required to normalize reaction conditions.

Fast photochemical oxidation of proteins (FPOP) is a mass spectrometry-based structural biology technique that probes the solvent-accessible surface area ...

Rapid Determination of Antibody-Antigen Affinity by Mass Photometry

Measurements of the specificity and affinity of antigen-antibody interactions are critically important for medical and research applications. In this protocol, we describe the implementation of a new single-molecule technique, mass photometry (MP), for this purpose. MP is a label- and immobilization-free technique that detects and quantifies molecular masses and populations of antibodies and antigen-antibody complexes on a single-molecule level. MP analyzes the antigen-antibody sample within minutes, allowing for the precise determination of the binding affinity and simultaneously providing information on the stoichiometry and the oligomeric state of the proteins. This is a simple and straightforward technique that requires only picomole quantities of protein and no expensive consumables. The same procedure can be used to study protein-protein binding for proteins with a molecular mass larger than 50 kDa. For multivalent protein interactions, the affinities of multiple binding sites can be obtained in a single measurement. However, the single-molecule mode of measurement and the lack of labeling imposes some experimental limitations. This method gives the best results when applied to measurements of sub-micromolar interaction affinities, antigens with a molecular mass of 20 kDa or larger, and relatively pure protein samples. We also describe the procedure for performing the required fitting and calculation steps using basic data analysis software.

We describe a single-molecule approach to antigen-antibody affinity measurements using mass photometry (MP). The MP-based protocol is fast, accurate, uses a very small amount of material, and does not require protein modification.

Measurements of the specificity and affinity of antigen-antibody interactions are critically important for medical and research applications. In this ...

Crystallization and Structural Determination of an Enzyme:Substrate Complex by Serial Crystallography in a Versatile Microfluidic Chip

The preparation of well diffracting crystals and their handling before their X-ray analysis are two critical steps of biocrystallographic studies. We describe a versatile microfluidic chip that enables the production of crystals by the efficient method of counter-diffusion. The convection-free environment provided by the microfluidic channels is ideal for crystal growth and useful to diffuse a substrate into the active site of the crystalline enzyme. Here we applied this approach to the CCA-adding enzyme of the psychrophilic bacterium Planococcus halocryophilus in the presented example. After crystallization and substrate diffusion/soaking, the crystal structure of the enzyme:substrate complex was determined at room temperature by serial crystallography and the analysis of multiple crystals directly inside the chip. The whole procedure preserves the genuine diffraction properties of the samples because it requires no crystal handling.

A versatile microfluidic device is described that enables the crystallization of an enzyme using the counter-diffusion method, the introduction of a substrate in the crystals by soaking, and the 3D structure determination of the enzyme:substrate complex by a serial analysis of crystals inside the chip at room temperature.

The preparation of well diffracting crystals and their handling before their X-ray analysis are two critical steps of biocrystallographic studies. We ...