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 ...

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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>Leptolyngbya 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 spectra</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>Synechocystis 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>&#946;-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 centri...

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. ...

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3.6K 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

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 ...

Isolation, Culture, and Functional Characterization of Adult Mouse Cardiomyoctyes

The use of primary cardiomyocytes (CMs) in culture has provided a powerful complement to murine models of heart disease in advancing our understanding of heart disease. In particular, the ability to study ion homeostasis, ion channel function, cellular excitability and excitation-contraction coupling and their alterations in diseased conditions and by disease-causing mutations have led to significant insights into cardiac diseases. Furthermore, the lack of an adequate immortalized cell line to mimic adult CMs, and the limitations of neonatal CMs (which lack many of the structural and functional biomechanics characteristic of adult CMs) in culture have hampered our understanding of the complex interplay between signaling pathways, ion channels and contractile properties in the adult heart strengthening the importance of studying adult isolated cardiomyocytes. Here, we present methods for the isolation, culture, manipulation of gene expression by adenoviral-expressed proteins, and subsequent functional analysis of cardiomyocytes from the adult mouse. The use of these techniques will help to develop mechanistic insight into signaling pathways that regulate cellular excitability, Ca2+ dynamics and contractility and provide a much more physiologically relevant characterization of cardiovascular disease.

Here we describe the isolation of adult mouse cardiomyoctyes using a Langendorff perfusion system. The resulting cells are Ca2+-tolerant, electrically quiescent and can be cultured and transfected with adeno- or lentiviruses to manipulate gene expression. Their functionality can also be analyzed using the MMSYS system and patch clamp techniques.

The use of primary cardiomyocytes (CMs) in culture has provided a powerful complement to murine models of heart disease in advancing our understanding of ...

En Face Detection of Nitric Oxide and Superoxide in Endothelial Layer of Intact Arteries

Endothelium-derived nitric oxide (NO) produced from endothelial NO-synthase (eNOS) is one of the most important vasoprotective molecules in cardiovascular physiology. Dysfunctional eNOS such as uncoupling of eNOS leads to decrease in NO bioavailability and increase in superoxide anion (O2.&#8722;) production, and in turn promotes cardiovascular diseases. Therefore, appropriate measurement of NO and O2.&#8722; levels in the endothelial cells are pivotal for research on cardiovascular diseases and complications. Because of the extremely labile nature of NO and O2.&#8722;, it is difficult to measure NO and O2.&#8722; directly in a blood vessel. Numerous methods have been developed to measure NO and O2.&#8722; production. It is, however, either insensitive, or non-specific, or technically demanding and requires special equipment. Here we describe an adaption of the fluorescence dye method for en face simultaneous detection and visualization of intracellular NO and O2.&#8722; using the cell permeable diaminofluorescein-2 diacetate (DAF-2DA) and dihydroethidium (DHE), respectively, in intact aortas of an obesity mouse model induced by high-fat-diet feeding. We could demonstrate decreased intracellular NO and enhanced O2.&#8722; levels in the freshly isolated intact aortas of obesity mouse as compared to the control lean mouse. We demonstrate that this method is an easy technique for direct detection and visualization of NO and O2.&#8722; in the intact blood vessels and can be widely applied for investigation of endothelial (dys)function under (physio)pathological conditions.

En Face Detection of Nitric Oxide and Superoxide in Endothelial Layer of Intact Arteries

In this article we describe an adapted relatively easy method using the fluorescence dye diaminofluorescein-2 diacetate (DAF-2DA) and dihydroethidium (DHE) for en face simultaneous detection and visualization of intracellular nitric oxide (NO) and superoxide anion (O2.&#8722;) respectively, in freshly isolated intact aortas of an obesity mouse model.

Endothelium-derived nitric oxide (NO) produced from endothelial NO-synthase (eNOS) is one of the most important vasoprotective molecules in cardiovascular ...

Isolation and Characterization of Microvesicles from Peripheral Blood

The release of extracellular vesicles (EVs) including small endosomal-derived exosomes (Exos, diameter &#60; 100 nm) and large plasma membrane-derived microvesicles (MVs, diameter &#62; 100 nm) is a fundamental cellular process that occurs in all living cells. These vesicles transport proteins, lipids and nucleic acids specific for their cell of origin and in vitro studies have highlighted their importance as mediators of intercellular communication. EVs have been successfully isolated from various body fluids and especially EVs in blood have been identified as promising biomarkers for cancer or infectious diseases. In order to allow the study of MV subpopulations in blood, we present a protocol for the standardized isolation and characterization of MVs from peripheral blood samples. MVs are pelleted from EDTA-anticoagulated plasma samples by differential centrifugation and typically possess a diameter of 100 - 600 nm. Due to their larger size, they can easily be studied by flow cytometry, a technique that is routinely used in clinical diagnostics and available in most laboratories. Several examples for quality control assays of the isolated MVs will be given and markers that can be used for the discrimination of different MV subpopulations in blood will be presented.

Extracellular vesicles present in blood have been suggested as novel biomarkers for various diseases. Here, we present a protocol for the isolation of large plasma membrane-derived microvesicles from peripheral blood samples and their subsequent analysis by conventional flow cytometry and Western Blotting.

The release of extracellular vesicles (EVs) including small endosomal-derived exosomes (Exos, diameter &#60; 100 nm) and large plasma membrane-derived ...

Isolation of Intact, Whole Mouse Mammary Glands for Analysis of Extracellular Matrix Expression and Gland Morphology

The goal of this procedure was to harvest the #4 abdominal mammary glands from female nulliparous mice in order to assess ECM expression and ductal architecture. Here, a small pocket below the skin was created using Mayo scissors, allowing separation of the glands within the subcutaneous tissue from the underlying peritoneum. Visualization of the glands was aided by the use of 3.5x-R surgical micro loupes. The pelt was inverted and pinned back allowing identification of the intact mammary fat pads. Each of the #4 abdominal glands was bluntly dissected by sliding the scalpel blade laterally between the subcutaneous layer and the glands. Immediately post-harvest, glands were placed in 10% neutral buffered formalin for subsequent tissue processing. Excision of the entire gland is advantageous because it primarily eliminates the risk of excluding important tissue-wide interactions between ductal epithelial cells and other microenvironmental cellular populations that could be missed in a partial biopsy. One drawback of the methodology is the use of serial sections from fixed tissues which limits analyses of ductal morphogenesis and protein expression to discrete locations within the gland. As such, changes in ductal architecture and protein expression in 3 dimensions (3D) is not readily obtainable. Overall, the technique is applicable to studies requiring whole intact murine mammary glands for downstream investigations such as developmental ductal morphogenesis or breast cancer.

Here, we present a protocol for the isolation of whole, intact mouse mammary glands to investigate extracellular matrix (ECM) expression and ductal morphology. Mouse #4 abdominal glands were extracted from 8-10 week old female nulliparous mice, fixed in neutral buffered formalin, sectioned and stained using immunohistochemistry for ECM proteins.

The goal of this procedure was to harvest the #4 abdominal mammary glands from female nulliparous mice in order to assess ECM expression and ductal ...

Visualization of DNA Compaction in Cyanobacteria by High-voltage Cryo-electron Tomography

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. DNA compaction is a dramatic cytoplasmic event recently found to occur in some cyanobacteria before cell division. However, due to the large cell size and the transient character, it is difficult to investigate the structure in detail. To overcome the difficulties, first, DNA compaction is reproducibly produced in the cyanobacterium Synechococcus elongatus PCC 7942 by synchronous culture using 12 h each light/dark cycle. Second, DNA compaction is monitored by fluorescence microscopy and captured by rapid freezing. Third, the detailed structure of DNA compacted cells is visualized in three dimensions (3D) by high voltage cryo-electron tomography. This set of methods is widely applicable to investigate transient structures in bacteria, e.g. cell division, chromosome segregation, phage infection etc., which are monitored by fluorescence microscopy and directly visualized by cryo-electron tomography at appropriate time points.

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. Synchronous cultivation, monitoring by fluorescence microscopy, rapid freezing, and high voltage cryo-electron tomography are used. A protocol for these methodologies is presented, and future applications and developments are discussed.

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. DNA compaction is a dramatic cytoplasmic event recently found to ...

Isolation of Intact Eyeball to Obtain Integral Ocular Surface Tissue for Histological Examination and Immunohistochemistry

Ocular surface (OS) consists of an epithelial sheet with three connected parts: palpebral conjunctiva, bulbar conjunctiva and corneal epithelium. Disruption of OS would lead to keratitis, conjunctivitis or both (keratoconjunctivitis). In experimental animal models with certain genetic modifications or artificial operations, it is useful to examine all parts of the OS epithelial sheet to evaluate relative pathogenetic changes of each part in parallel. However, dissection of OS tissue as a whole without distortion or damage has been challenging, primarily due to the softness and thinness of the OS affixed to physically separate yet movable eyelids and eyeball. Additionally, the deep eye socket formed by the hard skull/orbital bones is fully occupied by the eyeball leaving limited space for operating dissections. As a result, direct dissection of the eyeball with associated OS tissues from the facial side would often lead to tissue damages, especially the palpebral and bulbar conjunctiva. In this protocol, we described a method to remove the skull and orbital bones sequentially from a bisected mouse head, leaving eyelids, ocular surface, lens and retina altogether in one piece. The integrity of the OS sheet was well preserved and could be examined by histology or immunostaining in a single section.

This protocol describes a method for the isolation of the mouse eyeball with eyelid, ocular surface, anterior and posterior segments in relatively intact position.

Ocular surface (OS) consists of an epithelial sheet with three connected parts: palpebral conjunctiva, bulbar conjunctiva and corneal epithelium. ...

Isolation and Characterization of Adult Cardiac Fibroblasts and Myofibroblasts

Cardiac fibrosis in response to injury is a physiological response to wound healing. Efforts have been made to study and target fibroblast subtypes that mitigate fibrosis. However, fibroblast research has been hindered due to the lack of universally acceptable fibroblast markers to identify quiescent as well as activated fibroblasts. Fibroblasts are a heterogenous cell population, making them difficult to isolate and characterize. The presented protocol describes three different methods to enrich fibroblasts and myofibroblasts from uninjured and injured mouse hearts. Using a standard and reliable protocol to isolate fibroblasts will enable the study of their roles in homeostasis as well as fibrosis modulation.

Obtaining a pure population of fibroblasts is crucial to studying their role in wound repair and fibrosis. Described here is a detailed method to isolate fibroblasts and myofibroblasts from uninjured and injured mouse hearts followed by characterization of their purity and functionality by immunofluorescence, RTPCR, fluorescence-assisted cell sorting, and collagen gel contraction.

Cardiac fibrosis in response to injury is a physiological response to wound healing. Efforts have been made to study and target fibroblast subtypes that ...

Isolation and Characterization of Exosomes from Skeletal Muscle Fibroblasts

Exosomes are small extracellular vesicles released by virtually all cells and secreted in all biological fluids. Many methods have been developed for the isolation of these vesicles, including ultracentrifugation, ultrafiltration, and size exclusion chromatography. However, not all are suitable for large scale exosome purification and characterization. Outlined here is a protocol for establishing cultures of primary fibroblasts isolated from adult mouse skeletal muscles, followed by purification and characterization of exosomes from the culture media of these cells. The method is based on the use of sequential centrifugation steps followed by sucrose density gradients. Purity of the exosomal preparations is then validated by western blot analyses using a battery of canonical markers (i.e., Alix, CD9, and CD81). The protocol describes how to isolate and concentrate bioactive exosomes for electron microscopy, mass spectrometry, and uptake experiments for functional studies. It can easily be scaled up or down and adapted for exosome isolation from different cell types, tissues, and biological fluids.

This protocol illustrates the 1) the isolation and culture of primary fibroblasts from the adult mouse gastrocnemius muscle as well as 2) purification and characterization of exosomes using a differential ultracentrifugation method combined with sucrose density gradients followed by western blot analyses.

Exosomes are small extracellular vesicles released by virtually all cells and secreted in all biological fluids. Many methods have been developed for the ...

Isolation and Characterization of Mouse Primary Liver Sinusoidal Endothelial Cells

Liver sinusoidal endothelial cells (LSECs) are specialized endothelial cells located at the interface between the circulation and the liver parenchyma. LSECs have a distinct morphology characterized by the presence of fenestrae and the absence of basement membrane. LSECs play essential roles in many pathological disorders in the liver, including metabolic dysregulation, inflammation, fibrosis, angiogenesis, and carcinogenesis. However, little has been published about the isolation and characterization of the LSECs. Here, this protocol discusses the isolation of LSEC from both healthy and nonalcoholic fatty liver disease (NAFLD) mice. The protocol is based on collagenase perfusion of the mouse liver and magnetic beads positive selection of nonparenchymal cells to purify LSECs. This study characterizes LSECs using specific markers by flow cytometry and identifies the characteristic phenotypic features by scanning electron microscopy. LSECs isolated following this protocol can be used for functional studies, including adhesion and permeability assays, as well as downstream studies for a particular pathway of interest. In addition, these LSECs can be pooled or used individually, allowing multi-omics data generation including RNA-seq bulk or single cell, proteomic or phospho-proteomics, and Assay for Transposase-Accessible Chromatin using sequencing (ATAC-seq), among others. This protocol will be useful for investigators studying LSECs&#39; communication with other liver cells in health and disease and allow an in-depth understanding of the role of LSECs in the pathogenic mechanisms of acute and chronic liver injury.

Here we outline and demonstrate a protocol for primary mouse liver sinusoidal endothelial cell (LSEC) isolation. The protocol is based on liver collagenase perfusion, nonparenchymal cell purification by low-speed centrifugation, and CD146 magnetic bead selection. We also phenotype and characterize these isolated LSECs using flow cytometry and scanning electron microscopy.

Liver sinusoidal endothelial cells (LSECs) are specialized endothelial cells located at the interface between the circulation and the liver parenchyma. ...