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. Even though the analysis requires similar time and laboratory equipment as some of the previously published methods12,13,14,15,16, the advantage of this protocol is that no chlorophyll a is released into the extraction buffer and, thus, the phycobiliprotein measurement can be estimated with high precision.
The amount of cell suspension required for phycobiliprotein analysis can vary with the culture density. The sample volume of 1 mL is optimized for cultures with an OD730 of 1.5, a cell density of approximately 2 x 107 cells/mL, or for cultures with a phycobiliprotein content of approximately 20 mg/L. In the case of diluted cultures, harvest a larger culture volume. On the other hand, in the case of dense cultures, harvest a lower culture volume-in high-density cultures, there is a risk that phycobiliproteins partially remain in the cells after the extraction. Similarly, the amount of cell suspension required for the dry weight determination can vary with the culture density. The sampling volume of 15 mL is optimized for the cultures with an OD730 of 1.5 or with a cell density of approximately 2 x 107 cells/mL. With lower density cultures or with lower volumes, the measurement error can increase. On the contrary, bigger culture volumes or denser cultures provide better method resolutions.
Critical steps of the protocol are the cell homogenization and phycobiliprotein extraction that should provide both high yields and high specificity. The highest yields and purity of the extracts with simultaneous maximum protein preservation was achieved by freeze-drying the samples, homogenizing the dry cellular pellets by glass beads, and performing the phycobiliprotein extraction in PBS buffer (Figure 1). Since even a small contamination of the crude protein extract by chlorophyll a can lead to a phycobiliprotein content overestimation, it is necessary to avoid any chlorophyll a extraction by adding the PBS buffer. Chlorophyll a was detected in the crude extract when sonication was used for cell disruption. As shown in the inset of Figure 1, with repeating sonication cycles, the amount of chlorophyll a in the extract increased. On the other hand, the amount of extracted proteins (as detected by the absorbance at 280 nm) also increased after repeating sonication cycles. Therefore, the use of sonication (in combination with freezing-thawing cycles in liquid nitrogen) appears as a suitable method for total protein extraction (both free and membrane-bound proteins are released from the cells); however, it is not recommended for the purpose of phycobiliprotein spectrophotometric quantification. Chung et al. recommended the use of 1% streptomycin sulfate in order to eliminate membrane fragments with chlorophyll a16. Here, the use of streptomycin sulfate appeared not to be necessary since it did not lead to a reduction of chlorophyll a in the protein extract (Figure 7). Even though a single sonication cycle for 120 s did not disrupt the cells efficiently (methods B and C, Figure 1), other methods (e.g., method F as presented in Figure 1 or the methods presented in Figure 7B) confirmed previous findings of sonication being an efficient cell disruption method22,28. On the other hand, simple freezing-thawing cycles, also previously reported as an efficient method for phycobiliproteins extraction22,28, provided the lowest yields in the tests described here (method E, Figure 1). Cell homogenization (within the extraction buffer, 2 x 60 s) by zirconium beads was tested as less efficient than the 15-s homogenization of the freeze-dried cellular pellet (method D, Figure 1). The homogenization procedure within the extraction buffer was described previously with a longer homogenization time (10 min)29. We did not test the homogenization for longer than 2 min since the samples heated up significantly during each homogenization cycle. In the literature, other cell disruption methods were also described, including the utilization of a French press16 or griding13,15,30; however, such methods were not tested in this study.
The protein extraction was performed for 15 s (Figure 2) in PBS buffer. The buffer pH was 7.4, which was reported previously as optimal for phycobiliprotein extraction22. The addition of NaCl or KCl did not improve phycobiliprotein extraction efficiency (Figure 3), which is contradicting previous observations22. However, as indicated in Figure 3, the phosphate-buffered saline solution as used in this protocol contained 154 mM NaCl (in addition to 5.6 mM Na2HPO4 and 1 mM KH2PO4), which did not allow us to establish the NaCl-free PBS extraction buffer as a control. We also compared PBS buffer and sodium acetate buffer16 extraction efficiencies. The combination of sodium phosphate and potassium phosphate within the PBS buffer provided higher phycobiliproteins yields (Figure 3). This finding corresponded with the previous findings of Hemlata et al.22.
The phycobiliprotein extraction time was optimized for 1 h. Longer extraction did not lead to significant phycobiliprotein degradation or extraction improvement (Figure 4), which corresponded with the previous work of Lawrenz et al.28. An extraction time shorter than 1 h was not tested. Similarly, extraction for more than 4 h was not tested since it was found previously that the extracted phycobiliproteins in the phosphate buffer are stable for at least 48 h28.
We compared several equations of spectrophotometric phycocyanin and allophycocyanin quantification as described previously in the literature, by recalculating the content of both phycobiliproteins in the commercial standards with known protein concentrations (Figure 6). The best reconstruction of both pigments was achieved with the equations of Bennett and Bogorad12. This result was not specific to the phosphate extraction buffer since such a buffer has been used previously12,13,15. It was neither specific to the pigment standards since the difference between A615, A618, and A620 (wavelengths used for phycocyanin estimation in previous works) in the phycocyanin standard was 1% and the difference between A650 and A652 (wavelengths used previously for allophycocyanin estimation12,13,14,16) in the allophycocyanin standard was 3%. Similarly, the difference between A615 and A620 in the protein extract of Synechocystis was only 2%. Such small differences in the absorption spectra cannot result in a final pigment variation of up to 36% (Figure 6). Therefore, the differences between the individual equations were rather connected to variations in phycobiliprotein extinction coefficients of the specific organisms12,13,14,15,16,17. Interestingly, the equations of Chung et al. provided the lowest phycocyanin reconstruction (Figure 6) although the authors also used Synechocystis for their experiments16.
Instead of the determination of single wavelengths, it is recommended to measure a continuous spectrum of the phycobiliprotein extract between 280 - 720 nm, to estimate the presence of chlorophyll a in the extract, which could interfere with both the allophycocyanin and the phycocyanin absorption. Furthermore, by measuring the absorbance at 280 nm (A280), a purity of both phycocyanin (A615/A280 or A620/A280)21,22 and allophycocyanin (A652/A280)24 within the extract can be estimated (0.7: food grade, 3.9: reactive grade, &#62; 4.0: analytical grade)21.
As representative data, the concentration of both phycocyanin and allophycocyanin in Synechocystis was determined under increasing light intensity. Maintaining the culture density at a constant level within a turbidostat cultivation regime11,21 was essential for the direct comparative experiment. The phycocyanin and allophycocyanin concentrations of 2.4% - 10.2% and 0.6% - 1.7% of cellular dry weight, respectively, were similar as the previously reported values11,31,32. Also, the phycobiliprotein content per cell basis was similar as reported previously11,33. With the increasing light, the phycobiliproteins content decreased. Interestingly, even under the highest light intensity of 1100 &#181;mol(photons)/(m2&#183;s), the phycobiliproteins were not completely bleached.
Since the protocol requires only standard equipment and is relatively fast, it can be easily adopted in any laboratory in which phycobiliproteins are routinely analyzed.

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.

<table><tbody><tr><td>Synechocystis sp. PCC 6803</td><td>Institut Pasteur, Paris, France</td><td>6803</td><td>Cyanobacterium strain</td></tr><tr><td>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></tr><tr><td>Eppendorf safe-lock tubes&amp;nbsp;</td><td>Eppendorf, Hamburk, Germany</td><td>30120086</td><td>Safe-lock tubes 1.5 ml</td></tr><tr><td>VWR 80-Place Storage System</td><td>VWR International, Radnor, Pennsylvania, USA</td><td>30128-282</td><td>Holder for safe-lock tubes&amp;nbsp;</td></tr><tr><td>RAININ 100 &amp;micro;l -1000 &amp;micro;l&amp;nbsp;</td><td>Mettler-Toledo, Columbus, Ohio, USA</td><td>17014382</td><td>Pipette</td></tr><tr><td>GP-LTS-A-1000&amp;micro;L-/F-768/8</td><td>Mettler-Toledo, Columbus, Ohio, USA</td><td>30389272</td><td>Pipette tips</td></tr><tr><td>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></tr><tr><td>LCexv 4010</td><td>Liebherr, Bulle, Switzerland</td><td>9005382197172</td><td>Refrigerator and freezer -20&amp;nbsp;&amp;deg;C</td></tr><tr><td>Revco ExF -86&amp;deg;C Upright Ultra-Low Temperature Freezer</td><td>Thermo Fisher Scientific, Waltham, Massachusetts, USA</td><td>EXF24086V&amp;nbsp;</td><td>Freezer -80&amp;nbsp;&amp;deg;C</td></tr><tr><td>CoolSafe</td><td>LaboGene, Liller&amp;oslash;d, Denmark</td><td>7.001.000.615</td><td>Freeze dryer&amp;nbsp;</td></tr><tr><td>UV-2600</td><td>Shimadzu, Kyoto, Japan</td><td>UV-2600</td><td>Spectrophotometer&amp;nbsp;</td></tr><tr><td>Hellma absorption cuvettes, semi Micro</td><td>Sigma-Aldrich, St. Louis, Missouri, USA</td><td>Z600288&amp;nbsp;</td><td>VIS/UV-VIS semi-micro cuvettes 0.75-1.5 ml, spectral range 200-2500 nm&amp;nbsp;</td></tr><tr><td>Silamat S6</td><td>Ivoclar Vivadent, Schaan, Liechtenstein</td><td>602286WU</td><td>Homogenizer&amp;nbsp;</td></tr><tr><td>Solid-glass beads</td><td>Sigma-Aldrich, St. Louis, Missouri, USA</td><td>Z273627</td><td>Glass bead of the diameter 2&amp;nbsp;mm</td></tr><tr><td>CPA225D-0CE</td><td>Sartorius AG, G&amp;ouml;ttingen, Germany</td><td>SECURA225D-1OBR</td><td>Analytical balances</td></tr><tr><td>C-Phycocyanin from Spirulina sp.&amp;nbsp;</td><td>Sigma-Aldrich, St. Louis, Missouri, USA</td><td>P2172</td><td>Phycocyanin standard</td></tr><tr><td>Allophycocyanin</td><td>Sigma-Aldrich, St. Louis, Missouri, USA</td><td>A7472</td><td>Allophycocyanin standard</td></tr><tr><td>Bicinchoninic Acid Kit&amp;nbsp;</td><td>Sigma-Aldrich, St. Louis, Missouri, USA</td><td>BCA1, B9643</td><td>Complete kit for total proteins determination</td></tr><tr><td>AlgaeTron&amp;nbsp;</td><td>Photon System Instruments Ltd., Dr&amp;aacute;sov, Czech Republic</td><td>AG 130-ECO&amp;nbsp;</td><td>Cultivation chamber for E. flasks, with controllable light and atmosphere</td></tr><tr><td>Photobioreactor</td><td>Photon System Instruments Ltd., Dr&amp;aacute;sov, Czech Republic</td><td>FMT-150</td><td>Cultivation equipment for cyanobacteria and algae with completely controllable environment</td></tr><tr><td>Cellometer&amp;nbsp;</td><td>Nexcelom Bioscience, Lawrence, Massachusetts, USA</td><td>Auto M10</td><td>Cell counter</td></tr><tr><td>Corning 15 mL centrifuge tubes</td><td>Sigma-Aldrich, St. Louis, Missouri, USA</td><td>CLS430791&amp;nbsp;</td><td>15 ml Centrifuge tube for dry weigth sampling</td></tr><tr><td>Herasafe KS</td><td>Thermo Fisher Scientific, Waltham, Massachusetts, USA</td><td>51024579</td><td>Laminar flow hood</td></tr></tbody></table>

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 temperature for 5 min), the supernatant was discarded, and the samples were either stored at -80 &#176;C and freeze-dried as preparation for the individual steps of this protocol or stored at -20 &#176;C or -80 &#176;C for comparative extraction and quantifications analysis.
The tested extraction procedures included cell disruption by sonication, homogenization with zirconia beads within the PBS buffer, and freeze-thawing. The results of different extraction methods are summarized in Figure 1. The method as described within this protocol provided the highest phycobiliproteins yields together with the lowest chlorophyll a contamination in the extraction buffer. Chlorophyll a was detected in the extraction buffer when sonication was used for the cell disruption. Repeating cycles of freezing and thawing the samples in an ultrasound bath on ice resulted in higher protein yields in the extraction buffer. However, at the same time, the concentration of chlorophyll a in the extraction buffer increased (Figure 1 inset), which excluded a proper phycobiliprotein content quantification.
The optimal homogenization time was 15 s. Homogenization for both shorter (5 s) and longer (20 s) periods provided slightly lower phycobiliprotein yields, as shown in Figure 2. Homogenizing the cells with glass beads of a diameter of 2 mm resulted in the highest yields when compared with glass beads of a diameter of 0.5 mm, 1 mm, or 4 mm, with zirconia beads of a 0.5-mm diameter, or with sea sand grains. The PBS extraction buffer was tested as the optimal buffer for phycobiliproteins extraction (Figure 3). The addition of 100 mM NaCl or 150 mM KCl either to the PBS buffer or to water did not increase phycobiliprotein extraction efficiency, and the same went for using 20 mM sodium acetate buffer for extraction (Figure 3). The optimal phycobiliprotein extraction time was 60 minutes because, after more time (up to 240 minutes), the phycobiliprotein concentration in the extraction buffer did not change significantly (Figure 4).
We also tested the spectrophotometer linearity range for the phycobiliprotein measurement (Figure 5) and compared several equations of phycobiliprotein quantification, using both phycocyanin and allophycocyanin standards (Figure 6). The spectrophotometer used in this protocol showed a linear absorbance range between 0.1 - 3.0 (Figure 5). Since the spectra of the protein extract had its maximal absorption around 620 nm (A620), and at 652 nm, the absorption (A652) was only 33% of A620 (Figure 7), the spectrophotometer linearity for A652 (maximum allophycocyanin absorbance) was tested only up to A652 of 1.16. The equations (1) and (2) of Bennett and Bogorad12 provided the best reconstruction of both phycocyanin and allophycocyanin standards among all tested equations (Figure 6). Streptomycin sulfate, used in several previous studies for the elimination of membrane fragments containing chlorophyll a, was shown as not necessary for the extraction (Figure 7).
For the representative experiment, Synechocystis was cultivated in a photobioreactor25 in a turbidostat regime, where the cells were maintained in exponential growth phase within a defined range of optical density (measured at 680 nm, OD680) by a controlled dilution by a fresh cultivation medium (BG11 medium buffered with 17 mM HEPES)11,26. The cultures were cultivated at 32 &#176;C and the input air contained 0.5% CO2. The cultures were illuminated with a red light (&#955;max: 633 nm, &#955;1/2: 20 nm) of an intensity of 25 - 1100 &#181;mol(photons)/(m2&#183;s), together with a blue light (&#955;max: 445 nm, &#955;1/2: 20 nm) of an intensity of 25 &#181;mol(photons)/(m2&#183;s). The optical density of the culture suspension was measured by the photobioreactor base, and the range of OD680 was set to 0.52 - 0.58 (2 x 107 to 4 x 107 cells/mL). The cultures were cultivated under each specific light intensity for at least 24 hours to provide enough time for acclimation. After reaching growth stability, the cultures were sampled for dry weight (according to steps 2.1 - 2.8 of this protocol), cell count, and phycocyanin and allophycocyanin content. The results of the phycobiliprotein evaluation under increasing light intensities are shown in Figure 8. Synechocystis decreased the phycobiliprotein content from 12% of cellular dry weight (505 fg/cell) under the lowest irradiance to 3% of cellular dry weight (280 fg/cell) under the highest irradiance.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58076/58076fig1.jpg" /> 
Figure 1: Comparison of the extraction efficiency of the method described in this protocol with several reference methods. The concentration of phycocyanin (black bars) and allophycocyanin (white bars) in the crude protein extract was measured after extraction in the PBS buffer for 60 min. Method A is described within this protocol. Method B was modified as follows: after harvesting, the cells were centrifuged (15,000 x g at laboratory temperature for 5 min) and stored at -20 &#176;C overnight. The frozen samples were sonicated for 120 s at 4 &#176;C, and the phycobiliproteins were extracted in PBS buffer for 60 min. Method C was modified from method B by storing the pellets of harvested cells at -80 &#176;C for 30 min, freeze-drying the cellular pellets, and adding PBS buffer to the dry pellets. Method D was modified from Chung et al.16: after freeze-drying (same as in method C), 0.25 mL of Na-acetate with 1% streptomycin sulfate buffer (pH 5.5) was added to the dry cellular pellet, together with 0.25 mL of zirconium beads (with a diameter of 0.5 mm), and the samples were homogenized twice for 60 s on the homogenizer. After homogenization, another 0.5 mL of Na-acetate with 1% streptomycin sulfate buffer (pH 5.5) was added to the sample, the tubes were vortexed, and the proteins were extracted on ice for 30 min. Method E consisted of a single freezing-thawing cycle in PBS buffer and protein extraction for 24 h at 4 &#176;C. Method F (inset figure) consisted of one and six cycles of freezing the cells (initially, freeze-dried and resuspended in PBS buffer) in liquid nitrogen, followed by sonication on ice. Due to significant chlorophyll a presence in the crude protein extract, the phycobiliproteins were not quantified within method F. The depicted values represent averages from three technical replicates, with the error bars representing standard deviations. Pigment concentrations were calculated according to equations (1) and (2) of this protocol. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58076/58076fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58076/58076fig2.jpg" /> 
Figure 2: Effect of the homogenization time on the phycobiliprotein extraction efficiency. The lyophilized pellets of&#160;Synechocystis cells (according to steps 3.1 - 3.5 of this protocol) were disrupted with glass beads on homogenizer for 5 s, 15 s, and 20 s. The depicted values represent averages from three technical replicates, with the error bars representing standard deviations. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58076/58076fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58076/58076fig3.jpg" /> 
Figure 3: Phycobiliprotein extraction efficiency in various extraction buffers. The phycobiliproteins were extracted in the PBS buffer, in the PBS buffer with an addition of 100 mM NaCl or 150 mM KCl, in deionized water with 100 mM NaCl or 150 mM KCl, according to Hemlata and Fareha22, and in 20 mM sodium acetate according to Chung et al.16. The extraction was performed at 4 &#176;C according to steps 4.1 - 4.3 of this protocol. The depicted values represent averages from three technical replicates, with the error bars representing standard deviations. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58076/58076fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58076/58076fig4.jpg" /> 
Figure 4: Phycobiliprotein stability in the extraction buffer in time. After the extraction procedure, performed as described in steps 4.1 - 4.3 of this protocol, the phycobiliprotein extract in PBS buffer (pH 7.4) was stored at 4 &#176;C for up to 4 hours, with no significant phycocyanin (circles) and allophycocyanin (squares) degradation. The dashed line represents the linear fit of the time points calculated by the least squares method. The depicted values represent averages from three technical replicates, with the error bars representing standard deviations. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58076/58076fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/58076/58076fig5.jpg" /> 
Figure 5: Representative measurements of phycocyanin and allophycocyanin concentrations in Synechocystis. A dense culture suspension (phycocyanin: 456 &#181;g/mL, allophycocyanin: 106 &#181;g/mL) was gradually diluted up to a concentration of 19 &#181;g/mL for both phycocyanin and allophycocyanin. The depicted values represent averages from three technical replicates, with the error bars representing standard deviations. The dotted line represents the linear fit of the concentration points calculated by the least squares method. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58076/58076fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/58076/58076fig6.jpg" /> 
Figure 6: Estimation of phycocyanin and allophycocyanin concentrations in pigments standards. The content of phycocyanin and allophycocyanin in pigment standards was measured spectrophotometrically according to the equations of Bennett and Bogorad12, L&#252;der et al.13, Sampath-Wiley and Neefus15, Evans14, and Chung et al.16. Protein content in the standards (as necessary for phycobiliproteins determination) was quantified using a solution of bicinchoninic acid, sodium carbonate, sodium tartrate, and sodium bicarbonate in 0.1 N NaOH (with a final pH of 11.25), in reaction with 4% (w/v) copper(II) sulfate pentahydrate27, using bovine serum albumin as a protein standard. The equations of Bennett and Bogorad provided the highest reconstruction of both pigments: 96% of phycocyanin standard and 99% of allophycocyanin standard. The depicted values represent averages from three technical replicates, with the error bars representing standard deviations. The dashed line highlights the 100% point. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58076/58076fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/58076/58076fig7.jpg" /> 
Figure 7: The effect of streptomycin sulfate on the absorption spectrum of the crude protein extract in the PBS buffer. (A) The extraction procedure as described in steps 4.1 - 4.3 of this protocol or (B) using sonication as described in the legend of Figure 1 for Method F was performed in PBS buffer (circles) and in PBS buffer supplemented with 1% streptomycin sulfate (squares). The phycobiliprotein quantification was performed according to steps 5.1 - 5.4 of this protocol. The depicted values represent averages from three technical replicates, with the error bars representing standard deviations. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58076/58076fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/58076/58076fig8.jpg" /> 
Figure 8: Concentration of phycocyanin (circles) and allophycocyanin (squares) in Synechocystis cultivated under red-orange light of an intensity of 25 - 1100 &#181;mol(photons)/(m2&#183;s).&#160;Synechocystis cultures were cultivated in a photobioreactor at 32 &#176;C, with 0.5% CO2 in the input air, in BG11 cultivation medium supplemented with 17 mM HEPES in a turbidostat regime according to Zavrel et al.11. The phycobiliprotein concentrations were measured according to this protocol and the final phycobiliprotein content was normalized per cellular dry weight, as described in section 2 of this protocol. The dashed lines represent the power fit of the measured points, calculated by the least squares method. The depicted values represent averages from three technical replicates, with the error bars representing standard deviations.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58076/58076fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Nanyang Technological University

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JoVE Journal

Biochemistry

15.2K Views.  Czech Academy of Sciences. Hier presenteren we een protocol om te phycobiliprotein inhoud in de cyanobacterium Synechocystis een spectrofotometrische methode kwantitatief te bepalen. De extractiemethode werd ook met succes toegepast op andere cyanobacteriën en algen stammen; wegens verschillen in pigment absorptiespectra is het echter noodzakelijk is om de spectrofotometrische vergelijkingen voor elke soort afzonderlijk onderzoek.

Video: Spectrophotometric Determination of Phycobiliprotein Content in Cyanobacterium Synechocystis

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 ...

Isolating and Incorporating Light-Harvesting Antennas from Diatom Cyclotella Meneghiniana in Liposomes with Thylakoid Lipids

The photosynthetic performance of plants, algae and diatoms strongly depends on the fast and efficient regulation of the light harvesting and energy transfer processes in the thylakoid membrane of chloroplasts. The light harvesting antenna of diatoms, the so called fucoxanthin chlorophyll a/c binding proteins (FCP), are required for the light absorption and efficient transfer to the photosynthetic reaction centers as well as for photo-protection from excessive light. The switch between these two functions is a long-standing matter of research. Many of these studies have been carried out with FCP in detergent micelles. For interaction studies, the detergents have been removed, which led to an unspecific aggregation of FCP complexes. In this approach, it is hard to discriminate between artifacts and physiologically relevant data. Hence, more valuable information about FCP and other membrane bound light harvesting complexes can be obtained by studying protein-protein interactions, energy transfer and other spectroscopic features if they are embedded in their native lipid environment. The main advantage is that liposomes have a defined size and a defined lipid/protein ratio by which the extent of FCP clustering is controlled. Further, changes in the pH and ion composition that regulate light harvesting in vivo can easily be simulated. In comparison to the thylakoid membrane, the liposomes are more homogenous and less complex, which makes it easier to obtain and understand spectroscopic data. The protocol describes the procedure of FCP isolation and purification, liposome preparation, and incorporation of FCP into liposomes with natural lipid composition. Results from a typical application are given and discussed.

Isolating and Incorporating Light-Harvesting Antennas from Diatom Cyclotella Meneghiniana in Liposomes with Thylakoid Lipids

Here, we present a protocol to isolate fucoxanthin chlorophyll a/c binding proteins (FCP) from diatoms and incorporate them into liposomes with natural lipid compositions to study excitation energy transfer upon ion composition changes.

The photosynthetic performance of plants, algae and diatoms strongly depends on the fast and efficient regulation of the light harvesting and energy ...

Purification of Active Photosystem I-Light Harvesting Complex I from Plant Tissues

This method is used to isolate Photosystem I (PSI) together with the Light Harvesting Complex I (LHCI), its native antenna, from plants. PSI-LHCI is a large membrane protein complex coordinating hundreds of light harvesting and electron transport factors and is the most efficient light harvesting system found in nature. Photons absorbed by the four LHCA antenna proteins that make up LHCI are transferred through excitonic interaction to the PSI core reaction center and are used to facilitate light-driven charge separation across the thylakoid membrane, providing reducing power and energy for carbon fixation in photoautotrophic organisms. The high quantum efficiency of PSI makes this complex an excellent model to study light-driven energy transfer. In this protocol, plant tissue is mechanically homogenized, and the chloroplasts are separated from the bulk cellular debris by filtration and centrifugation. The isolated chloroplasts are then osmotically lysed, and the thylakoid membranes are recovered via centrifugation and solubilized using the detergent n-dodecyl-beta-maltoside. The solubilized material is loaded onto an anion exchange column to collect most of the chlorophyll-containing complexes. Larger complexes are precipitated from the solution, resuspended in a small volume, and loaded on sucrose gradients to separate the major chlorophyll-containing complexes. The resulting sucrose gradient fractions are characterized to identify the band of interest containing PSI-LHCI. This protocol is highly similar to the protocol used in the crystallization of plant PSI-LHCI with some simplifications and relies on methods developed over the years in the lab of Nathan Nelson.

This protocol describes the isolation of Photosystem I (PSI) - Light Harvesting Complex I (LHCI) from plant tissues. PSI along with PSII is responsible for the conversion of light to chemical energy in oxygenic photoautotrophs and has a quantum efficiency of ~1, making it a target for studying light-driven energy transfer.

This method is used to isolate Photosystem I (PSI) together with the Light Harvesting Complex I (LHCI), its native antenna, from plants. PSI-LHCI is a ...

Photobleaching Enables Super-resolution Imaging of the FtsZ Ring in the Cyanobacterium Prochlorococcus

Super-resolution microscopy has been widely used to study protein interactions and subcellular structures in many organisms. In photosynthetic organisms, however, the lateral resolution of super-resolution imaging is only ~100 nm. The low resolution is mainly due to the high autofluorescence background of photosynthetic cells caused by high-intensity lasers that are required for super-resolution imaging, such as stochastic optical reconstruction microscopy (STORM). Here, we describe a photobleaching-assisted STORM method which was developed recently for imaging the marine picocyanobacterium Prochlorococcus. After photobleaching, the autofluorescence of Prochlorococcus is effectively reduced so that STORM can be performed with a lateral resolution of ~10 nm. Using this method, we acquire the in vivo three-dimensional (3-D) organization of the FtsZ protein and characterize four different FtsZ ring morphologies during the cell cycle of Prochlorococcus. The method we describe here might be adopted for the super-resolution imaging of other photosynthetic organisms.

Photobleaching Enables Super-resolution Imaging of the FtsZ Ring in the Cyanobacterium Prochlorococcus

Here we describe a photobleaching method to reduce the autofluorescence of cyanobacteria. After photobleaching, stochastic optical reconstruction microscopy is used to obtain three-dimensional super-resolution images of the cyanobacterial FtsZ ring.

Super-resolution microscopy has been widely used to study protein interactions and subcellular structures in many organisms. In photosynthetic organisms, ...

Immunology and Infection

Protocol for Co-Culturing Sucrose-Secreting Cyanobacteria with Fungal Partners

Assembly and Quantification of Co-Cultures Combining Heterotrophic Yeast with Phototrophic Sugar-Secreting Cyanobacteria

With the increasing demand for sustainable biotechnologies, mixed consortia containing a phototrophic microbe and heterotrophic partner species are being explored as a method for solar-driven bioproduction. One approach involves the use of CO2-fixing cyanobacteria that secrete organic carbon to support the metabolism of a co-cultivated heterotroph, which in turn transforms the carbon into higher-value goods or services. In this protocol, a technical description to assist the experimentalist in the establishment of a co-culture combining a sucrose-secreting cyanobacterial strain with a fungal partner(s), as represented by model yeast species, is provided. The protocol describes the key prerequisites for co-culture establishment: Defining the media composition, monitoring the growth characteristics of individual partners, and the analysis of mixed cultures with multiple species combined in the same growth vessel. Basic laboratory techniques for co-culture monitoring, including microscopy, cell counter, and single-cell flow cytometry, are summarized, and examples of nonproprietary software to use for data analysis of raw flow cytometry standard (FCS) files in line with FAIR (Findable, Accessible, Interoperable, Reusable) principles are provided. Finally, commentary on the bottlenecks and pitfalls frequently encountered when attempting to establish a co-culture with sugar-secreting cyanobacteria and a novel heterotrophic partner is included. This protocol provides a resource for researchers attempting to establish a new pair of co-cultured microbes that includes a cyanobacterium and a heterotrophic microbe.

This protocol provides a comprehensive guideline for the setup and quantitative monitoring of co-cultures, including photoautotrophic sugar-secreting cyanobacteria and heterotrophic yeasts.

With the increasing demand for sustainable biotechnologies, mixed consortia containing a phototrophic microbe and heterotrophic partner species are being ...

Bioengineering

Measuring Photophysiology of Attached Stage of Colacium sp. by a Cuvette-Type Fast Repetition Rate Fluorometer

Fast repetition rate fluorometer (FRRf) is a beneficial method for measuring photosystem II (PSII) photophysiology and primary productivity. Although FRRf can measure PSII absorption cross-section (&#963;PSII), maximum photochemical efficiency (Fv/Fm), effective photochemical efficiency (Fq&#8242;/Fm&#8242;), and non-photochemical quenching (NPQNSV) for various eukaryotic algae and cyanobacteria, almost all FRRf studies to date have focused on phytoplankton. Here, the protocol describes how to measure PSII photophysiology of an epizoic alga Colacium sp.&#160;Ehrenberg 1834 (Euglenophyta), in its attached stage (attached to zooplankton), using cuvette-type FRRf. First, we estimated the effects of substrate zooplankton (Scapholeberis mucronata O.F. M&#252;ller 1776, Cladocera, Daphniidae) on baseline fluorescence and &#963;PSII, Fv/Fm, Fq&#8242;/Fm&#8242;, and NPQNSV of planktonic Colacium sp. To validate this methodology, we recorded photophysiology measurements of attached Colacium sp. on S. mucronata and compared these results with its planktonic stage. Representative results showed how the protocol could determine the effects of calcium (Ca) and manganese (Mn) on Colacium sp. photophysiology and identify the various effects of Mn enrichment between attached and planktonic stages. Finally, we discuss the adaptability of this protocol to other periphytic algae.

Measuring Photophysiology of Attached Stage of Colacium sp. by a Cuvette-Type Fast Repetition Rate Fluorometer

Fast repetition rate fluorometer (FRRf) is a beneficial method for measuring photosystem II photophysiology and primary productivity. Here we describe a protocol to measure PSII photophysiology of epizoic alga, Colacium sp. on substrate zooplankton using cuvette-type FRRf.

Fast repetition rate fluorometer (FRRf) is a beneficial method for measuring photosystem II (PSII) photophysiology and primary productivity. Although FRRf ...

Biology

Isolation and Characterization of Intact Phycobilisome in Cyanobacteria

In cyanobacteria, phycobilisome is a vital antenna protein complex that harvests light and transfers energy to photosystem I and II for photochemistry. Studying the structure and composition of phycobilisome is of great interest to scientists because it reveals the evolution and divergence of photosynthesis in cyanobacteria. This protocol provides a detailed and optimized method to break cyanobacterial cells at low cost by a bead-beater efficiently. The intact phycobilisome can then be isolated from the cell extract by sucrose gradient ultracentrifugation. This method has demonstrated being suitable for both model and non-model cyanobacteria with different cell types. A step-by-step procedure is also provided to confirm the integrity and property of phycobiliproteins by 77K fluorescence spectroscopy and SDS-PAGE stained by zinc sulfate and Coomassie Blue. The isolated phycobilisome can also be subjected to further structural and compositional analyses. Overall, this protocol provides a helpful starting guide that allows researchers unfamiliar with cyanobacteria to quickly isolate and characterize intact phycobilisome.

The current protocol details the isolation of phycobilisomes from cyanobacteria by centrifugation through a discontinuous sucrose density gradient. The fractions of intact phycobilisomes are confirmed by 77K fluorescent emission spectrum and SDS-PAGE analysis. The resulting phycobilisome fractions are suitable for negative staining of TEM and mass spectrometry analysis.

In cyanobacteria, phycobilisome is a vital antenna protein complex that harvests light and transfers energy to photosystem I and II for photochemistry. ...

Natural Transformation, Protein Expression, and Cryoconservation of the Filamentous Cyanobacterium Phormidium lacuna

Cyanobacteria are the focus of basic research and biotechnological projects in which solar energy is utilized for biomass production. Phormidium lacuna is a newly isolated filamentous cyanobacterium. This paper describes how new filamentous cyanobacteria can be isolated from marine rockpools. It also describes how DNA can be extracted from filaments and how the genomes can be sequenced. Although transformation is established for many single-celled species, it is less frequently reported for filamentous cyanobacteria. A simplified method for the natural transformation of P. lacuna is described here. P. lacuna is the only member of the order Oscillatoriales for which natural transformation is established. This paper also shows how natural transformation is used to express superfolder green fluorescent protein (sfGFP). An endogenous cpcB promoter induced approximately 5 times stronger expression than cpc560, A2813, or psbA2 promoters from Synechocystis sp. PCC6803. Further, a method for the cryopreservation of P. lacuna and Synechocystis sp. CPP 6803 was established, and methods for assessing motility in a liquid medium and on agar and plastic surfaces are described.

Natural Transformation, Protein Expression, and Cryoconservation of the Filamentous Cyanobacterium Phormidium lacuna

Phormidium lacuna is a filamentous cyanobacterium that was isolated from marine rockpools. This article describes the isolation of filaments from natural sources, DNA extraction, genome sequencing, natural transformation, expression of sfGFP, cryoconservation, and motility methods.

Cyanobacteria are the focus of basic research and biotechnological projects in which solar energy is utilized for biomass production. Phormidium lacuna is ...

Autofluorescence Imaging to Evaluate Red Algae Physiology

Red algae (Rhodophyta) contain phycobiliproteins and colonize habitats with dim light, however some (e.g., some Chroothece species) can also develop in full sunshine. Most rhodophytes are red, however some can appear bluish, depending on the proportion of blue and red biliproteins (phycocyanin and phycoerythrin). Different phycobiliproteins can capture light at diverse wavelengths and transmit it to chlorophyll a, which makes photosynthesis under very different light conditions possible. These pigments respond to habitat changes in light, and their autofluorescence can help to study biological processes. Using Chroothece mobilis as a model organism and the spectral lambda scan mode in a confocal microscope, the adaptation of photosynthetic pigments to different monochromatic lights was studied at the cellular level to guess the species' optimal growth conditions. The results showed that, even when the studied strain was isolated from a cave, it adapted to both dim and medium light intensities. The presented method is especially useful for studying photosynthetic organisms that do not grow or grow very slowly under laboratory conditions, which is usually the case for those living in extreme habitats.

The present protocol describes step-by-step autofluorescence imaging and evaluation of phycobiliprotein changes in red algae based on spectral analysis. This is a label-free and non-destructive method to evaluate cellular adaptation to extreme habitats, when only scarce material is available and cells grow slowly, or not at all, under laboratory conditions.

Red algae (Rhodophyta) contain phycobiliproteins and colonize habitats with dim light, however some (e.g., some Chroothece species) can also develop in ...

Immunocytochemical Visualization of Proteins from Cyanobacterial Cells with High Autofluorescence of Phycoerythrin and Phycourobilin

Presented is a protocol for visualizing and quantifying a specific protein in cells at the cellular level for the marine cyanobacterium Crocosphaera watsonii, a crucial primary producer and nitrogen fixer in oligotrophic oceans. One of the challenges for marine autotrophic N2 fixers (diazotrophs) is distinguishing probe-derived fluorescence signals from autofluorescence. C. watsonii was selected to represent chlorophyll-, phycoerythrin- and phycourobilin-containing cyanobacteria. The protocol allows for simple and semi-quantitative visualization of proteins in C. watsonii at a single-cell level, enabling investigation of protein production under different environmental conditions to evaluate the metabolic activities of the target cyanobacteria. Furthermore, the fixation and permeabilization methods are optimized to enhance the fluorescence signals from target proteins to distinguish them from autofluorescence, especially from phycoerythrin and phycourobilin. The enhanced signal can be visualized using confocal or widefield fluorescence microscopy. Additionally, fluorescence intensity was semi-quantified using Fiji software. This single-cell analysis workflow allows the evaluation of cell-to-cell variations of specific protein content. The protocol can be performed in any life science laboratory as it requires only standard equipment and can also be easily adapted to other phycoerythrin-containing cyanobacterial cells.

This protocol outlines the visualization and quantification of a particular protein within cells at the cellular level for the phycoerythrin-containing cyanobacterium, Crocosphaera watsonii.

Presented is a protocol for visualizing and quantifying a specific protein in cells at the cellular level for the marine cyanobacterium Crocosphaera ...

Spectrophotometric Determination of Phycobiliprotein Content in Cyanobacterium Synechocystis

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