Name: isolation and characterization of intact phycobilisome in cyanobacteria
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The authors thank Technology Commons, College of Life Science, National Taiwan University for the convenient use of the ultracentrifuge. The cyanobacterial strains Synechocystis sp. PCC 6803 and Leptolyngbya sp. JSC-1 was gifted from Dr. Chu, Hsiu-An at Academia Sinica, Taiwan, and Dr. Donald A. Bryant at Pennsylvania State University, USA, respectively. This work was funded by the Ministry of Science and Technology (Taiwan) (109-2636-B-002-013- and 110-2628-B-002-065-) and the Ministry of Education (Taiwan) Yushan Young Scholar Program (109V1102 and 110V1102).

The Synechocystis sp.&#160;PCC 6803, the model glucose-tolerant strain, was obtained from Dr. Chu, Hsiu-An at Academia Sinica, Taiwan. Leptolyngbya sp. JSC-1, the non-model filamentous,was obtained from Dr. Donald A. Bryant at Pennsylvania State University, USA.
1. Cell culture and harvesting
<ol>
	<li>Inoculate Syn6803 or JSC-1&#160;cells using a metallic inoculation loop into a 100 mL conical flask containing 50 mL of B-HEPES medium28. Culture the cells at 30 &#176;C under 50 &#181;mol photons m-2 s-1 (white-light LED) in 1% (v/v) CO2 with constant stirring by a magnetic stirrer (120 rpm) until the culture reaches mid-exponential growth phase (OD750 = 0.6-0.8). 
	NOTE: Harvest the cell culture when the optical density of the culture at 750 nm (OD750) reaches 0.6-0.8 by transferring to a new flask. Optical density was routinely used to estimate microbial cell density in liquid cultures29. The wavelengths from 720-750 nm were used in various studies for measuring cyanobacterial growth30,31,32. In this protocol, OD750 is used because JSC-1 can produce pigments that absorb far-red light7. Alternatively, chlorophyll concentration was also used to measure the growth of cyanobacteria33.</li>
	<li>Transfer the cell culture (OD750 ~0.8) into a 1L conical flask and dilute it to OD750 = 0.2 in 500 mL of B-HEPES medium. Grow the culture until the late exponential growth phase (OD750 = 0.8-1.0) in the same condition mentioned above (step 1.1) and then harvest the culture by centrifugation.</li>
	<li>Centrifuge the culture at 10,000 x g for 20 min at room temperature and completely discard the supernatant. 
	&#8203;NOTE: The cells pellets can be stored at -80 &#176;C for up to 6 months.</li>
</ol>
2. Cells lysis
<ol>
	<li>Suspend the cell pellets and wash twice with 0.75 M K-phosphate buffer, pH 7. 
	NOTE: To make 0.75 M K-phosphate buffer at pH 7, prepare 1.5 M of K2HPO4 and 1.5 M of KH2PO4 separately. Mix 615 mL of 1.5 M K2HPO4 and 385 ml of 1.5M KH2PO4 to get 1.5 M K-phosphate buffer at pH 7. Simply dilute it with the same amount of ddH2O to make 0.75 M K-phosphate buffer at pH 7.</li>
	<li>Pellet the cells in 50 mL centrifuge tubes by centrifugation at 10,000 x g for 20 min at room temperature. Discard the supernatant and resuspend the cells with 0.75 M K-phosphate buffer. Measure the wet weight of the pellet by an electronic balance and resuspend 1 g wet weight of the pellet in 5 mL of the buffer. 
	NOTE: Wet weight of a pellet was measured by subtracting the weight of an empty centrifuge tube from the weight of the centrifuge tube containing the cell pellet.</li>
	<li>Add 1 mL of cell suspension and 0.4-0.6 g of 0.1 mm glass beads (see Table of Materials) into a 2 mL screw cap vial.</li>
	<li>Break the cells for 30 s by a bead-beater (see Table of Materials). Allow the tubes to cool in a room temperature water bath for 2 min and repeat the breaking cycle 5 times.</li>
	<li>After lysis, centrifuge the vials at 500 x g for 10 s at room temperature. Collect the supernatant with a pipette into a 15 mL centrifuge tube. Wash the beads with 0.5 mL of 0.75 M K-phosphate buffer one time, then transfer to the same centrifuge tube.</li>
	<li>Add Triton X-100 (2% w/v, final concentration) to the lysed cell suspension and incubate on a rocking shaker (40 rpm) at room temperature until the solution becomes homogenous (~20 min).</li>
	<li>Centrifuge the tubes at 17,210 g for 20 min to remove the intact cells and large cell debris. Store the supernatant without the upper Triton micelle layer at room temperature for up to 1 h. 
	&#8203;NOTE: The supernatant contains two separated layers. The upper hydrophobic Triton X layer contains chlorophyll-binding proteins and hydrophobic rod-shaped phycobilisomes6. The lower aqueous layer contains water-soluble PBS.</li>
</ol>
3. Preparation of the sucrose gradient buffers and PBS isolation
<ol>
	<li>Make four concentrations of sucrose (2.0 M, 1.0 M, 0.75 M and 0.5 M) in 0.75 M K-phosphate buffer at pH 7. 
	NOTE: It is suggested to use 1.5 M K-phosphate buffer at pH 7 to prepare the sucrose gradient because adding sucrose dilutes the concentration of K-phosphate buffer. After sucrose is fully dissolved, add ddH2O to adjust the concentration to 0.75 M, pH 7.</li>
	<li>Place 2.8 mL of 2.0 M sucrose buffer at the bottom of the 40 mL centrifuge tube and overlay with three layers of sucrose solutions (8 mL of 1.0 M; 12 mL of 0.75 M and 11 mL of 0.5 M for 40 mL centrifuge tube), and finally PBS-containing the supernatant fraction (3.0 mL) (Figure 1A). 
	NOTE: Carefully add the sucrose solution and allow sucrose to drop very slowly inside of the tube. Hold the pipette tip just above the surface of the solution in the tube while loading the solution. Sucrose layers can be observed when they are layered slowly.</li>
	<li>Centrifuge the resulting gradients at 125,800 x g for ~16h-20 h at 25 &#176;C.NOTE: An ultracentrifuge (see Table of Materials) is required for this step.</li>
	<li>Upon ultra-centrifugation, condense the fractionated PBS and phycobiliproteins between the layers. Blue bands in Syn6803 (purple bands showed in JSC-1) were observed in the interfaces (Figure 1D).</li>
	<li>Slowly collect the fractions with a pipette from the top of the sucrose gradients. Remove the sucrose by repeatedly concentrating (3,500 x g for 20 min) and diluting the fractions with 0.75 M K-phosphate buffer in membrane centrifugal filter units (10 K molecular weight cut-off, see Table of Materials).</li>
</ol>
4. Measurement of PBS fluorescence at 77K
NOTE: A fluorimeter equipped with a liquid nitrogen container (see Table of Materials) is used to measure fluorescence spectra at 77K.
<ol>
	<li>Dilute the concentrated PBS sample with 0.75 M K-phosphate buffer to gain at least 500 &#181;L. 
	NOTE: The concentrations of phycocyanin of samples were ~4.2 &#181;g mL-1 based on the formula: (OD615 0.474 x OD652)/5.34 [mg/mL]34.</li>
	<li>Add 500 &#181;L of the PBS sample to a transparent glass tube and freeze the tube in liquid nitrogen until it is completely frozen. Move the frozen tube to a transparent Dewar container (see Table of Materials) pre-filled with liquid nitrogen. 
	NOTE: The inner diameter of the glass tube is 3 mm in this protocol. Thin glass tubes were used to minimize re-absorption of short-wavelength emission in the sample.</li>
	<li>Choose the excitation wavelengths for phycoerythrin and phycocyanin to be 550 nm and 580 nm, respectively. 
	&#8203;NOTE: Fluorescence emissions were recorded from 560-800 nm (for 550 nm excitation) or 600-800 nm (for 580 nm excitation).</li>
</ol>
5. SDS-PAGE analysis of PBS
<ol>
	<li>Buffer exchange the concentrated PBS samples in 0.75 M of K-phosphate with 50 mM of Tris buffer (pH 8.0) in membrane centrifugal filter units (3K molecular weight cut-off, see Table of Materials).</li>
	<li>Mix 50 &#181;L of PBS solution with 10 &#181;L of 6x SDS loading buffer [300 mM Tris pH 6.8, 12% (w/v) Sodium dodecyl sulfate, 0.06% (w/v) Bromphenol blue, 50% (v/v) Glycerol, 6% (v/v) &#946;-Mercaptoethanol] (see Table of Materials) in a microcentrifuge tube and incubate at 95 &#176;C for 10 min in a heat block.</li>
	<li>Separate the protein samples by SDS-PAGE (8%-20% (w/v) polyacrylamide gel) and continue with zinc and Coomassie staining. The detailed procedure has been described in Reference8.</li>
	<li>For zinc staining, incubate the gel in 50 mM ZnSO4 solution for 10 min at room temperature, wash the gel with distilled water, and visualize zinc-induced fluorescence under UV irradiation (312 nm).</li>
	<li>For Coomassie blue staining, incubate the gel in Coomassie Blue staining buffer [0.25% (w/v) of Coomassie R-250, 10% (v/v) of Acetic acid, 50% (v/v) of Methanol, see Table of Materials] for 1 h at room temperature. Wash the gel with distilled water twice to remove the residual staining buffer and incubate the gel with destaining buffer [10% (v/v) Acetic acid, 30% (v/v) Methanol] on a rocking shaker (40 rpm) at room temperature overnight. Visualize the fully-destained gel with a digital camera or a scanner.</li>
</ol>

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structural+organisation+of+phycobilisomes+from+Synechocystis+sp.+strain+PCC6803+and+their+interaction+with+the+membrane.">Structural organisation of phycobilisomes from Synechocystis sp. strain PCC6803 and their interaction with the membrane.</a> Biochimica et Biophysica Acta (BBA) - Bioenergetics. 1787 (4), 272-279 (2009).</li><li>Kondo, K., Ochiai, Y., Katayama, M., Ikeuchi, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Membrane-associated+CpcG2-phycobilisome+in+Synechocystis:+A+new+photosystem+I+antenna.">The Membrane-associated CpcG2-phycobilisome in Synechocystis: A new photosystem I antenna.</a> Plant Physiology. 144 (2), 1200-1210 (2007).</li></ol>

The authors declare no competing interest.

This protocol describes a simple and standard method for isolating intact PBS in two types of cyanobacteria, unicellular model Syn6803,&#160;and filamentous non-model JSC-1. The critical steps of the protocol are cell homogenization and ultracentrifugation on a discontinuous density gradient of sucrose. Generally, the disruption of filamentous cells is more complicated than unicellular ones. Increasing the amount of the starting material (the wet weight of the cell pellet) and the repetition of bead-beating were helpful to increase the yield of PBS from filamentous cyanobacterial cells. For the filamentous cyanobacterium, JSC-1, three times more cells were used than unicellular one, Syn6803. In addition, to completely break the filamentous cyanobacterium, it requires bead-beating more times than the unicellular cyanobacteria to observe the dark bluish-purple color in the extraction buffer.
The duration of bead-beating is not suggested to exceed 30 s since the samples are heated up during each bead-beating cycle. The tubes are recommended to cool at room temperature on a bench (or in a water bath) between each cycle, as the intact PBS is readily dissociated at low temperature25. Therefore, every procedure in this protocol is suggested to be performed at room temperature. Many research articles have described different cell disruption methods, including a high-pressure homogenizer (French-Press) or a sonicator6,8,20. A bead mill homogenizer is suggested because it is cheaper, safer, easier to handle, faster, more accessible, and more efficient in processing multiple samples simultaneously.
Phycobilisomes show three different shapes, hemidiscoidal, bundle-shaped, and rod-shaped5. The intact phycobilisomes isolated in Figure 2 belong to the hemidiscoidal shape as discussed in earlier publications7,13,36. The upper fraction that represents dissociated PBS, however, can also contain rod-shaped PBS. Syn6803 and JSC-1 encode a specific rod-core linker CpcL, which suggests the presence of rod-shaped PBS in addition to the hemidiscoidal PBS6,37. Rod-shaped PBS has been directly identified in Syn6803 but not in JSC-17,13,37. On the other hand, JSC-1 remodels its PBS by performing type III chromatic acclimation in green/red light and far-red light photoacclimation in red/far-red light6,7. All these diversities indicate that there may be other types of PBS in the sucrose gradients, in addition to the lowest intact hemidiscoidal PBS fraction. For example, two types of PBS are present at the same time when Synechococcus sp. PCC 7335 and JSC-1 acclimates to far-red light8,13. It is suggested that the person isolating PBS from an uncharacterized cyanobacterial strain should be more cautious in analyzing each fraction in the sucrose gradient to identify any possible types of PBS.
Overall, this protocol provides a cheap, simple, and fast method for cell disruption. It has also been demonstrated that this method is reliable to isolate PBS from different cyanobacteria with different cell types and PBS compositions.

Phycobilisome (PBS) is a huge water-soluble pigment-protein complex that attaches to the cytoplasmic side of the photosystems in the thylakoid membranes of cyanobacteria1. PBS is primarily composed of colored phycobiliproteins and colorless linker proteins1,2. The phycobiliproteins can be divided into four major groups: phycoerythrin, phycoerythrocyanin, phycocyanin, and allophycocyanin3. The four major groups absorb different wavelengths of light energy in the range of 490-650 nm, which chlorophylls absorbed inefficiently3. The PBS can serve as a light-harvesting antenna for collecting light energy and delivering it to Photosystem II and I4.
The structure and composition of PBS vary from species to species. Collectively, three shapes of phycobilisome (hemidiscodial, bundle-shaped, and rod-shaped) have been identified in different cyanobacterial species5. Even in the same species, the composition of PBS changes in response to the environment, such as light quality and nutrient depletion6,7,8,9,10,11. Therefore, the experimental procedure to isolate PBS from cyanobacteria has been instrumental in studying PBS12. Over several decades, many different protocols have isolated PBS and analyzed its structure, composition, and function6,7,8,12,13,14,15,16,17. The wide variety of methods for PBS isolation indeed provides flexibility in isolating the complex in different species with different reagents and instruments. However, it also makes choosing a suitable protocol more difficult for scientists unfamiliar with cyanobacteria and PBS. Therefore, a generalized and straightforward protocol is developed in this work for those interested in starting PBS isolation from cyanobacteria.
The methods for isolating PBS from previous publications are summarized here. Since PBS is a water-soluble protein complex and is readily dissociated, a high ionic strength phosphate buffer is required to stabilize PBS during extraction18. Several research articles that describe methods for the isolation of PBS from cyanobacterium have been published in the past. Most of the methods require a high concentration of phosphate buffer8,14,15,18,19. However, the procedures for mechanical disrupting of the cells vary, such as glass beads-assisted extraction, sonication20, and French press6,8,14. Different phycobiliproteins can be obtained by precipitation with ammonium sulfate20 and purified by HPLC21 or a chromatographic column22. On the other hand, intact PBS can be easily isolated by sucrose density gradient ultracentrifugation6,8,15.
In this protocol, one model cyanobacterium and one non-model cyanobacterium were used as the materials for PBS isolation. They are model unicellular&#160;glucose-tolerant Synechocystis sp.&#160;PCC 6803 (hereafter Syn6803) and non-model filamentous Leptolyngbya sp. JSC-1 (hereafter JSC-1), respectively7,23,24. The protocol begins by disruption of the unicellular and filamentous cyanobacteria in a high-ionic-strength phosphate buffer. After lysis, the supernatants are collected by centrifugation and then treated with a nonionic detergent (Triton X-100) to solubilize the water-soluble proteins from the thylakoid membranes. The total water-soluble proteins are applied to a discontinuous sucrose density gradient to fractionate the PBS. The discontinuous sucrose gradient in this protocol consists of four sucrose solutions and partitions the intact PBS in the lowest fractions of sucrose layer25. The integrity of PBS can be analyzed by SDS-PAGE, zinc-staining, and 77K fluorescence spectroscopy6,7,8,26. This method is suitable for scientists who aim to isolate intact PBS from cyanobacteria and study its spectral, structural, and compositional properties.
There are several advantages of this protocol. (1) This method is standardized and can be used for isolating intact PBS from both unicellular and filamentous cyanobacteria. Most of the articles describe the method that applied in either one type of cyanobacteria4,7,8,12,13,14,16,18. (2) This method is performed at room temperature, as PBS dissociates at low temperature19,27. (3) This method describes using a bead-beater to disrupt the cells; therefore, it is cheaper and safer than high-pressured French press and possible hearing damage from sonicator in other methods8,13,14,20. (4) This method isolates intact PBS by sucrose gradient ultra-centrifugation. In this way, intact PBS with different sizes and partially dissociated PBS can be separated based on sucrose concentration.

<table><tbody><tr><td>0.1 mm glass beads</td><td>BioSpec</td><td>11079101</td><td>for PBS extraction</td></tr><tr><td>13 mL centrifugation tube</td><td>Hitachi</td><td>13PA</td><td>ultracentrifugation</td></tr><tr><td>40 mL centrifugation tube</td><td>Hitachi</td><td>40PA</td><td>ultracentrifugation</td></tr><tr><td>Acetic acid</td><td>Merck</td><td>8.1875.2500</td><td>for Coomassie Blue staining</td></tr><tr><td>B-HEPES medium</td><td></td><td></td><td>A modified cyanobacterial medium from BG-11 medium</td></tr><tr><td>Brilliant Blue R-250</td><td>Sigma</td><td>B-0149</td><td>for Coomassie Blue staining</td></tr><tr><td>Bromophenol blue</td><td>Wako pure chemical industries</td><td>2-291</td><td>protein loading buffer</td></tr><tr><td>Electronic balance</td><td>Radwag</td><td>WLC 2/A2/C/2</td><td>for the wet weight measurement of cell pellets</td></tr><tr><td>Fluorescence spectrophotometer</td><td>Hitachi</td><td>F-7000</td><td>Spectrophotometer</td></tr><tr><td>Glycerol</td><td>BioShop</td><td>Gly001.500</td><td>protein loading buffer</td></tr><tr><td>High-Speed refrigerated centrifuge</td><td>Hitachi</td><td>CR22N</td><td>for buffer exchange</td></tr><tr><td>&lt;em&gt;Leptolyngbya &lt;/em&gt;sp. JSC-1</td><td></td><td></td><td>from Dr. Donald A. Bryant at Pennsylvania State University, USA.</td></tr><tr><td>Low temperature measurement accessory</td><td>Hitachi</td><td>5J0-0112</td><td>The accessory includes a transparent Dewar container for 77K fluorescence spectroscopy.</td></tr><tr><td>Methanol</td><td>Merck</td><td>1.07018,2511</td><td>for Coomassie Blue staining</td></tr><tr><td>Microcentrifuge</td><td>Thermo Fisher</td><td>Pico 21</td><td>for PBS extraction</td></tr><tr><td>Mini-Beadbeater-16</td><td>BioSpec</td><td>Model 607</td><td>for PBS extraction</td></tr><tr><td>Potassium phosphate dibasic</td><td>PanReac AppliChem</td><td>121512.121</td><td>for PBS extraction</td></tr><tr><td>Potassium phosphate monobasic</td><td>PanReac AppliChem</td><td>141509.121</td><td>for PBS extraction</td></tr><tr><td>Screw cap vial</td><td>BioSpec</td><td>10832</td><td>for PBS extraction</td></tr><tr><td>SmartView Pro Imager</td><td>Major Science</td><td>UVCI-2300</td><td>for Znic staining signal detection</td></tr><tr><td>Sodium dodecyl sulfate</td><td>Zymeset</td><td>BSD101</td><td>protein loading buffer</td></tr><tr><td>Sucrose</td><td>Zymeset</td><td>BSU101</td><td>for PBS isolation</td></tr><tr><td>&lt;em&gt;Synechocystis &lt;/em&gt;sp. PCC 6803</td><td></td><td></td><td>glucose-tolerant strain from Dr. Chu, Hsiu-An at Academia Sinica, Taiwan</td></tr><tr><td>Tris</td><td>BioShop</td><td>TRS 011.1</td><td>protein loading buffer</td></tr><tr><td>Triton X-100</td><td>BioShop</td><td>TRX 506.500</td><td>for PBS extraction</td></tr><tr><td>Ultra 10 K membrane centrifugal filter</td><td>Millipore</td><td>UFC901024</td><td>for buffer exchange</td></tr><tr><td>Ultra 3 K membrane centrifugal filter</td><td>Millipore</td><td>UFC500324</td><td>for buffer exchange</td></tr><tr><td>Ultracentrifuge</td><td>Hitachi</td><td>CP80WX</td><td>ultracentrifugation</td></tr><tr><td>UV/Vis spectrophotometer</td><td>Agilent</td><td>Cary 60</td><td>Spectrophotometer</td></tr><tr><td>Zinc sulfate</td><td>PanReac AppliChem</td><td>131787.121</td><td>for Znic staining</td></tr><tr><td>&amp;beta;-Mercaptoethanol</td><td>BioBasic</td><td>MB0338</td><td>protein loading buffer</td></tr></tbody></table>

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 Syn6803 and JSC-1 cells were cultivated in conical flasks with constant stirring in B-HEPES medium at 30 &#176;C, under a LED white light (50 &#181;mol photons m-2s-1) in a growth chamber filled with 1% (v/v) CO2. At the exponential growth phase (OD750 = ~0.5), the cells were subcultured into fresh medium with a final optical density OD750 = ~0.2. After reaching the late exponential growth phase (OD750 = 0.6-0.8), the cultures were collected and centrifuged (10,000 x g at room temperature for 20 min). The supernatant was discarded, and the cell pellets were stored at -80 &#176;C until the next step was available.
To break the cells, the cells were first thawed at room temperature. The cells were disrupted by bead-beating with 0.1 mm glass beads within the 0.75 M K-phosphate buffer. Complete cell lysis shows the supernatant in dark blue-green color before centrifugation (Figure 1B). After solubilization by Triton X-100 and centrifugation to remove the insoluble membranes and cell debris, the supernatant was loaded on the top of sucrose gradient solution in a 40 mL centrifuge tube (Figure 1C). After 18 h of centrifugation, the intact PBS moved to the gradient&#39;s center as a clear blue band for Syn6803 and a purple band for JSC-1. The difference in their color results from the different compositions of PBS in Syn6803 and JSC-1. The PBS from Syn6803 carries phycocyanin35, whereas the PBS of JSC-1 carries phycocyanin and phycoerythrin7. The result of sucrose density-gradient separation of isolated PBS shows several fractions of dissociated phycobiliproteins, and the lowest fraction represents the intact PBS (Figure 1D).
The absorption spectra of the different fractions of PBS isolated from JSC-1 and Syn6803 are compared in Figure&#160;2A. The detailed procedure of absorption spectra has been described in the articles7,8. The absorption spectra from the dissociated PBS and intact PBS of JSC-1 and Syn6803 have the same peak at 622 nm, corresponding to the phycocyanin. And the phycoerythrin absorption peak at 571 nm is found in both dissociated PBS and intact PBS of JSC-1. Allophycocyanin shows a minor peak with a shoulder at 670 nm in the intact PBS fractions of JSC-1 and Syn6803. The spectral differences indicate that the rod (phycocyanin) to core (allophycocyanin) ratio is higher in the dissociated PBS than in the intact PBS fraction, as the spectral properties of phycobiliproteins have been determined previously5. Since the dissociated PBS and the intact PBS fractions show similar absorption spectra and fluorescent emission features at room temperature, 77K fluorescence spectroscopy was then used to analyze the different fractions isolated from the sucrose gradient. 77K fluorescent emission spectra of the dissociated PBS and the intact PBS fractions were excited by 580 nm. The emission spectra of the dissociated PBS have one strong peak at ~650 nm in Syn6803 and JSC-1, representing the emission from phycocyanin (Figure 2B). The fluorescent emission spectra of the intact PBS show two fluorescent emission maximal peaks: the weak one at 651 nm and the strong one at 684 nm, revealing the energy harvested by phycocyanin was transferred efficiently to allophycocyanin and terminal emitters (ApcD and ApcE) in the core9 (Figure 2C). These features demonstrate that the lowest fractions in the sucrose gradient contain intact PBS.
The dissociated PBS and the intact PBS were separated by SDS-PAGE on an 8%-20% (w/v) SDS polyacrylamide linear gradient gel. The &#945; and &#946; subunits of phycocyanin and allophycocyanin were visible on the gel without staining (Figure 3A). The gel was subsequently stained with 10 mM ZnSO4 for 10 min. The chromophore-containing phycobiliproteins were observed by the zinc staining during UV irradiation8. Phycobiliprotein subunits (14-21 kDa) are strongly fluorescent, and the ApcE (arrow) showed weak fluorescence (Figure 3B). Further, Coomassie Blue staining shows the different protein composition between the dissociated PBS fractions and the intact PBS fractions (Figure 3C). The upper fractions contain other non-PBS water-soluble proteins as more proteins are stained than in intact PBS fractions. The intact PBS in Syn6803 shows a prominent ApcE band (arrow) lacking in the dissociated PBS fractions (Figure 3C).
The volume ratio between the PBS extract and the sucrose gradient is critical for obtaining sharp bands in the sucrose gradient after ultracentrifugation. The suggested ratio of solubilized PBS extracts to sucrose gradient solution is 1:10. Based on previous experience, making the sucrose gradient in 40 mL centrifuge tubes shows a higher resolution than the sucrose gradient made in the 13 mL tubes. The discontinuous sucrose gradient diffuses over time because of vibration on the bench. Therefore, the 13 mL centrifuge tube can be a good choice for handling lots of samples simultaneously. To demonstrate that, the sucrose gradients in 13 mL centrifuge tubes at the ratios as 1:3 and 1:10 have been prepared between the crude extracts of Syn6803 or JSC-1 to the sucrose gradient solution (Figure 4A). After ultracentrifugation, the sucrose gradient prepared at ratio 1:3 displays wider bands than the gradient made at ratio 1:10 (Figure 4B). The lowest fractions of each gradient contain intact PBS, confirmed by 77K fluorescent emission spectroscopy.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63272/63272fig01.jpg" /> 
Figure 1: Disruption of Syn6803 and JSC-1 cells and sucrose gradient ultracentrifugation. (A) Schematic illustration of preparing the discontinued sucrose gradient in a 40 mL centrifuge tube.&#160;(B)&#160;The screw-cap vials were filled with 0.4 g of glass beads and 1 mL of cell suspension of Syn6803 or JSC-1, and then the cells were lysed by a bead-beater. (C) The supernatant of Triton X-100 solubilized extract was layered on the top of a sucrose density gradient. (D)&#160;After centrifugation for 18 h, the PBS fractions were visualized as clear blue bands in the gradient in Syn6803 and as purple bands in JSC-1. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/63272/63272fig01large.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/63272/63272fig02.jpg" /> 
Figure 2: Spectroscopic characterization of different fractions of sucrose gradients. (A) Absorption spectra of dissociated PBS and intact PBS fractions from JSC-1 and Syn6803.&#160;Fluorescence emission spectra were measured at 77 K using the excitation wavelength at 580 nm for (B) the dissociated PBS fractions and (C) the intact PBS fractions of Syn6803 and JSC-1. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/63272/63272fig02large.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/63272/63272fig03.jpg" /> 
Figure 3: SDS-PAGE analysis of PBS isolated from sucrose gradients. The dissociated and the intact PBS fractions were quantified (20 &#181;g protein per lane) and loaded onto an 8%-20% (w/v) gradient polyacrylamide gel. After gel electrophoresis, the gel was visualized by zinc staining and Coomassie Blue staining, subsequently. (A)&#160;The color of phycobiliproteins is visible on the gel without staining. (B) Zinc-stained gel shows the fluorescence of phycobiliproteins. (C) The gel stained with Coomassie Blue show phycobiliproteins and other proteins. The arrows indicate the positions of the ApcE. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/63272/63272fig03large.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/63272/63272fig04.jpg" /> 
Figure 4: The sucrose gradient made in 13 mL centrifuge tubes. The ratios between the crude extract and the sucrose gradient solution are marked on the top of the images. (A) The supernatant of Triton X-100 solubilized extracts of Syn6803 and JSC-1 was layered over sucrose density gradients with different ratios of the solubilized extracts to sucrose gradient solution. (B)&#160;After centrifugation for 18 h, the PBS fractions in Syn6803 and JSC-1 were observed in the sucrose gradients. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/63272/63272fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Isolation and Characterization of Intact Phycobilisome in Cyanobacteria at JoVE.com

Isolation and Characterization of Intact Phycobilisome in Cyanobacteria

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. ...

isolation-characterization-intact-phycobilisome

Universidad San Sebastian

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JoVE Journal

Biology

3.8K Views.  National Taiwan University. 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.

Video: Isolation and Characterization of Intact Phycobilisome in Cyanobacteria

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 ...

Biochemistry

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.

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.

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

Looking Outwards: Isolation of Cyanobacterial Released Carbohydrate Polymers and Proteins

Cyanobacteria can actively secrete a wide range of biomolecules into the extracellular environment, such as heteropolysaccharides and proteins. The identification and characterization of these biomolecules can improve knowledge about their secretion pathways and help to manipulate them. Furthermore, some of these biomolecules are also interesting in terms of biotechnological applications. Described here are two protocols for easy and rapid isolation of cyanobacterial released carbohydrate polymers and proteins. The method for isolation of released carbohydrate polymers is based on conventional precipitation techniques of polysaccharides in aqueous solutions using organic solvents. This method preserves the characteristics of the polymer and simultaneously avoids the presence of contaminants from cell debris and culture medium. At the end of the process, the lyophilized polymer is ready to be used or characterized or can be subjected to further rounds of purification, depending on the final intended use. Regarding the isolation of the cyanobacterial exoproteome, the technique is based on the concentration of the cell-free medium after removal of the major contaminants by centrifugation and filtration. This strategy allows for reliable isolation of proteins that reach the extracellular milieu via membrane transporters or outer membrane vesicles. These proteins can be subsequently identified using standard mass spectrometry techniques. The protocols presented here can be applied not only to a wide range of cyanobacteria, but also to other bacterial strains. Furthermore, these procedures can be easily tailored according to the final use of the products, purity degree required, and bacterial strain.

Here, protocols for the isolation of cyanobacterial released carbohydrate polymers and isolation of their exoproteomes are described. Both procedures embody key steps to obtain polymers or proteins with high purity degrees that can be used for further analysis or applications. They can also be easily adapted according to specific user needs.

Cyanobacteria can actively secrete a wide range of biomolecules into the extracellular environment, such as heteropolysaccharides and proteins. The ...

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 ...

Isolation and Characterization of RNA-Containing Exosomes

The field of exosome research is rapidly expanding, with a dramatic increase in publications in recent years. These small vesicles (30-100 nm) of endocytic origin were first proposed to function as a way for reticulocytes to eradicate the transferrin receptor while maturing into erythrocytes1, and were later named exosomes. Exosomes are formed by inward budding of late endosomes, producing multivesicular bodies (MVBs), and are released into the environment by fusion of the MVBs with the plasma membrane2. Since the first discovery of exosomes, a wide range of cells have been shown to release these vesicles. Exosomes have also been detected in several biological fluids, including plasma, nasal lavage fluid, saliva and breast milk3-6. Furthermore, it has been demonstrated that the content and function of exosomes depends on the originating cell and the conditions under which they are produced. A variety of functions have been demonstrated for exosomes, such as induction of tolerance against allergen7,8, eradication of established tumors in mice9, inhibition and activation of natural killer cells10-12, promotion of differentiation into T regulatory cells13, stimulation of T cell proliferation14 and induction of T cell apoptosis15. Year 2007 we demonstrated that exosomes released from mast cells contain messenger RNA (mRNA) and microRNA (miRNA), and that the RNA can be shuttled from one cell to another via exosomes. In the recipient cells, the mRNA shuttled by exosomes was shown to be translated into protein, suggesting a regulatory function of the transferred RNA16. Further, we have also shown that exosomes derived from cells grown under oxidative stress can induce tolerance against further stress in recipient cells and thus suggest a biological function of the exosomal shuttle RNA17. Cell culture media and biological fluids contain a mixture of vesicles and shed fragments. A high quality isolation method for exosomes, followed by characterization and identification of the exosomes and their content, is therefore crucial to distinguish exosomes from other vesicles and particles. Here, we present a method for the isolation of exosomes from both cell culture medium and body fluids. This isolation method is based on repeated centrifugation and filtration steps, followed by a final ultracentrifugation step in which the exosomes are pelleted. Important methods to identify the exosomes and characterize the exosomal morphology and protein content are highlighted, including electron microscopy, flow cytometry and Western blot. The purification of the total exosomal RNA is based on spin column chromatography and the exosomal RNA yield and size distribution is analyzed using a Bioanalyzer.

This paper demonstrates methods for the isolation, purification and detection of exosomes, as well as techniques for analysis of their molecular content. These methods are adaptable for exosome isolation from both cell culture media and biological fluids, and can beyond analysis of molecular content also be useful in functional studies.

The field of exosome research is rapidly expanding, with a dramatic increase in publications in recent years. These small vesicles (30-100 nm) of ...

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

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 ...

Isolation and Characterization of Cyanobacterial Extracellular Vesicles

Cyanobacteria are a diverse group of photosynthetic, Gram-negative bacteria that play critical roles in global ecosystems and serve as essential biotechnology models. Recent work has demonstrated that both marine and freshwater cyanobacteria produce extracellular vesicles -&#160;small membrane-bound structures released from the outer surface of the microbes. While vesicles likely contribute to diverse biological processes, their specific functional roles in cyanobacterial biology remain largely unknown. To encourage and advance research in this area, a detailed protocol is presented for isolating, concentrating, and purifying cyanobacterial extracellular vesicles. The current work discusses methodologies that have successfully isolated vesicles from large cultures of Prochlorococcus, Synechococcus, and Synechocystis. Methods for quantifying and characterizing vesicle samples from these strains are presented. Approaches for isolating vesicles from aquatic field samples are also described. Finally, typical challenges encountered with cyanobacterial vesicle purification, methodological considerations for different downstream applications, and the trade-offs between approaches are also discussed.

The present protocol providesdetailed descriptions for isolation, concentration,and characterization of extracellular vesicles from cyanobacterial cultures. Approaches for purifying vesicles from cultures at different scales, trade-offs among methodologies, and considerations for working with field samples are also discussed.

Cyanobacteria are a diverse group of photosynthetic, Gram-negative bacteria that play critical roles in global ecosystems and serve as essential ...

Plastoglobule Lipid Droplet Isolation from Plant Leaf Tissue and Cyanobacteria

Plastoglobule lipid droplets are a dynamic sub-compartment of plant chloroplasts and cyanobacteria. Found ubiquitously among photosynthetic species, they are believed to serve a central role in the adaptation and remodeling of the thylakoid membrane under rapidly changing environmental conditions. The capacity to isolate plastoglobules of high purity has greatly facilitated their study through proteomic, lipidomic, and other methodologies. With plastoglobules of high purity and yield, it is possible to investigate their lipid and protein composition, enzymatic activity, and protein topology, among other possible molecular characteristics. This article presents a rapid and effective protocol for the isolation of plastoglobules from chloroplasts of plant leaf tissue and presents methodological variations for the isolation of plastoglobules and related lipid droplet structures from maize leaves, the desiccated leaf tissue of the resurrection plant, Eragrostis nindensis, and the cyanobacterium, Synechocystis sp. PCC 6803. Isolation relies on the low density of these lipid-rich particles, which facilitates their purification by sucrose density flotation. This methodology will prove valuable in the study of plastoglobules from diverse species.

A fast and efficient protocol is presented for the isolation of plastoglobule lipid droplets associated with various photosynthetic organisms. The successful preparation of isolated plastoglobules is a crucial first step that precedes detailed molecular investigations such as proteomic and lipidomic analyses.

Plastoglobule lipid droplets are a dynamic sub-compartment of plant chloroplasts and cyanobacteria. Found ubiquitously among photosynthetic species, they ...

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 ...

Isolation and Characterization of Intact Phycobilisome in Cyanobacteria

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

Spectrophotometric Determination of Phycobiliprotein Content in Cyanobacterium Synechocystis

Looking Outwards: Isolation of Cyanobacterial Released Carbohydrate Polymers and Proteins

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

Isolation and Characterization of RNA-Containing Exosomes

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

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

Isolation and Characterization of Cyanobacterial Extracellular Vesicles

Plastoglobule Lipid Droplet Isolation from Plant Leaf Tissue and Cyanobacteria

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