The authors thank Daiying Xu for her technical assistance and comments on the manuscript. This study is supported by grants from the National Natural Science Foundation of China (Project number 41476147) and the Research Grants Council of the Hong Kong Special Administrative Region, China (project numbers 689813 and 16103414).

1. Sample Preparation and Fixation
<ol>
	<li>Inoculate 1 mL of axenic Prochlorococcus MED4 to 5 mL of the seawater-based Pro99 medium12. Grow Prochlorococcus cultures at 23 &#176;C under the light with an intensity of 35 &#181;mol photons/m2s. Five days later, collect 1 mL of culture into a 1.5-mL tube. 
	NOTE: Five days after the inoculation, Prochlorococcus MED4 will reach the late log phase, with approximately 108 cell/mL, which is appropriate for STORM imaging.</li>
	<li>Add 100 &#181;L of formaldehyde freshly prepared from 25% paraformaldehyde (PFA) (w/v) and 2 &#181;L of 50% glutaraldehyde into the 1.5-mL tube to fix the culture. Incubate the sample in the dark at room temperature for 20 min. 
	NOTE: The sample was kept in the dark for fixation because glutaraldehyde is light sensitive.</li>
	<li>Spin down the sample at 13,500 x g for 1 min; then, remove the supernatant and resuspend the cells in 100 &#181;L of the Pro99 medium12. Store the sample at 4 &#176;C until immunostaining.</li>
</ol>
2. Precoating of the Coverslip with Polystyrene Beads
NOTE: Polystyrene beads are considered as the fiducial marker for drift correction.
<ol>
	<li>Vortex the original vial of polystyrene beads and prepare a 1:20,000 dilution with 50% ethanol as working slurry.</li>
	<li>Turn on the hot plate, set the temperature to 120 &#176;C, and place coverslips onto the hot plate.</li>
	<li>Load 100 &#181;L of working slurry onto each coverslip. Incubate the coverslip on the hot plate for 10 min and, then, carefully transfer the coverslips to Petri dishes for storage at room temperature. 
	NOTE: The side with the beads attached to it should face up, and the cells should be attached on the same side. The beads are immobilized on the coverslips and used for correcting the sample-drifting in real-time during STORM imaging. After immobilizing the beads, the coverslips cannot be flamed.</li>
</ol>
3. Coating of Poly-L-lysine onto the Bead-coated Coverslip
NOTE: This is done for the immobilization of cyanobacterial cells.
<ol>
	<li>Add 100 &#181;L of poly-L-lysine (1 mg/mL) to the center of the coverslip with the bead-coated side facing up. Let it sit for 30 min at room temperature.</li>
	<li>Use a pipette to carefully aspire the unattached poly-L-lysine and transfer it to a 1.5-mL tube. Save the poly-L-lysine at 4 &#176;C for reuse.</li>
	<li>Rinse the coverslip with 10 mL of ultra-filtered water in a 60-mm Petri dish and then briefly dry the coverslip on a paper towel.</li>
	<li>Store the coverslip in a Petri dish (100 mm) for later usage. To prevent the attachment of the coverslip to the dish, line the Petri dish with a layer of parafilm. 
	NOTE: The coverslip, precoated with beads and poly-L-lysine, can be stored at room temperature for at least half a year. Therefore, preparing a batch of coverslips in advance is highly recommended.</li>
</ol>
4. Immobilization of Cells on the Coverslip
<ol>
	<li>Transfer a precoated coverslip to a new Petri dish with parafilm at the bottom, which will be referred to as the staining dish henceforth. Load 100 &#181;L of the fixed sample on the coverslip and let it sit for 30 min to allow cell attachment.</li>
	<li>Transfer the coverslip to a well of a 12-well plate, which will be referred to as the washing dish henceforth. Add 1 mL of phosphate-buffered saline (PBS) (10 mM Na2HPO4, 1.8 mM KH2PO4, 137 mM NaCl, and 2.7 mM KCl, pH 7.4) to the well to wash the coverslip. Remove the PBS buffer and add a new PBS buffer to wash the coverslip again.</li>
</ol>
5. Permeabilization of Cyanobacterial Cells
<ol>
	<li>Remove the PBS buffer using a pipette. Add 1 mL of freshly prepared permeabilization buffer to the well of the washing dish, which contains 0.05% non-ionic detergent-100 (v/v), 10 mM EDTA, 10 mM Tris (pH 8.0), and 0.2 mg/mL lysozyme.</li>
	<li>Incubate the washing dish at 37 &#176;C for 20 min. Then, remove the permeabilization buffer.</li>
	<li>Add 1 mL of PBS buffer and place the washing dish on a shaker which gently shakes the washing dish for 5 min. Remove the PBS buffer. Repeat this step 3x to wash the coverslip.</li>
</ol>
6. Photobleaching of the Chlorophyll Pigments in a Blocking Step
<ol>
	<li>Transfer the coverslip to the staining dish. Add 50 &#181;L of blocking buffer, which contains 0.2% (v/v) non-ionic detergent-100 and 3% (v/v) goat serum, on the top of the coverslip.</li>
	<li>Place the whole staining plate on ice and move it under the xenon light source for photobleaching for at least 60 min, with a light intensity at 1,800 &#181;mol photons/m2&#183;s.</li>
	<li>After photobleaching and blocking, carefully remove the blocking buffer by adsorbing it with the edge of a paper towel. 
	NOTE: The intensity of xenon light is so high that a pair of proper sunglasses is needed for eye protection. Meanwhile, wrap the light source and the plate using aluminum foil to avoid the leaking of light, which may cause eye damage. Check the coverslip every 15 min to make sure that the coverslip remains moist. If the coverslip is dry, add 50 &#181;L of blocking buffer.</li>
</ol>
7. Antibody Binding
<ol>
	<li>Dissolve anti-Anabaena FtsZ antibody in 200 &#181;L of water according to the manufacturer&#8217;s instructions.</li>
	<li>Dilute 1 &#181;L of anti-FtsZ antibody into 99 &#181;L of blocking buffer. Add 50 &#181;L of diluted primary antibody on the coverslip and incubate it at room temperature for 30 min.</li>
	<li>Transfer the coverslip to the washing dish. Add 1 mL of PBS buffer and place the washing dish on a shake. Gently shake the washing dish for 5 min. Repeat this step 3x to wash the coverslip.</li>
	<li>Transfer the coverslip to the staining dish. Dilute 1 &#181;L of a secondary antibody into 500 &#181;L of blocking buffer. Add 50 &#181;L of diluted secondary antibody onto the coverslip and incubate it for 30 min in the dark at room temperature.</li>
	<li>Transfer the coverslip to the washing dish. Wash it with 1 mL of PBS buffer on a shaker. Gently shake the washing dish for 5 min. Wrap the washing dish with aluminum paper to make it light-proof. Repeat this step 3x.</li>
	<li>Add 500 &#181;L of formaldehyde freshly prepared from 4% paraformaldehyde (PFA) (w/v) to the well. Incubate the coverslip for 15 min in the dark at room temperature.</li>
	<li>Remove the formaldehyde and repeat the washing step 3x.</li>
	<li>Store the coverslip in PBS buffer at 4 &#176;C in the dark until imaging.</li>
</ol>
8. Preparation of the STORM Imaging Buffer
<ol>
	<li>Prepare 1 mL of imaging buffer according to Table 1, immediately before the STORM imaging. 
	NOTE: As the shelf life of the imaging buffer is about 2 h, prepare it fresh, right before usage. Avoid vortexing after the addition of glucose oxidase and catalase. 
	CAUTION: Methyl viologen is poisonous material. Please handle it with particular care. Cyclooctatetraene is classified as a carcinogen Cat. 1 chemical. Please avoid inhalation and skin contact and make the cyclooctatetraene stock solution and imaging buffer in a chemical hood.</li>
</ol>
9. Image Acquisition of STORM Data
<ol>
	<li>Load the coverslip in the loading chamber (Figure 1). Load the freshly prepared imaging buffer gently into the chamber to avoid washing off the cyanobacterial cells. Place a rectangular coverslip on the top of the imaging buffer to prevent its reaction with oxygen in the air. Avoid trapping air bubbles underneath the coverslip.</li>
	<li>Turn on the camera, the LED light, and the laser. Open STORM software for image acquisition, centroid position determination, and sample drifting correction13.</li>
	<li>Add half a drop of immersion oil on top of the lens. Load the chamber and make sure the lens of the objective makes contact with the coverslip.</li>
	<li>Examine the signal with a 750-nm laser. 
	NOTE: Always include a second antibody-minus-staining negative control to ensure that the autofluorescence is diminished.</li>
	<li>Identify a sample area that contains both cells and fiducial markers. Start the software for sample drifting correction. 
	NOTE: In general, the ideal sample area contains 10 - 30 cells. Meanwhile, the cells should be separated well to avoid any overlapping cells.</li>
	<li>Acquire one wide-field image as a reference, with the camera electron multiplication (EM) gain at 300 and an exposure time of 30 ms (Figure 2A).</li>
	<li>Increase the 750-nm excitation laser intensity to a higher power, approximately 4.5 kW/cm2. Once the fluorophores have transitioned into a sparse blinking pattern, acquire one super-resolution image by collecting 10,000 frames at 33 Hz (Figure 2B). 
	NOTE: If the fluorophores are not isolated, wait for a couple of minutes until more fluorophores are switched to the dark state and the isolated single molecules flicker sequentially. The number of frames described here is suitable for our sample and needs to be optimized for other targeted structures.</li>
</ol>
10. Reconstruction of Super-resolution Images from Raw Data
<ol>
	<li>Use a plugin called QuickPALM (Table of Materials)14 in ImageJ to reconstruct a 3-D color super-resolution image. 
	NOTE: A preliminary chromatic image (Figure 2C) along with a color-coded scale bar (Figure 2C) will appear. The color represents the depth on the z-axis.</li>
	<li>To demonstrate the 3-D structure in a video, construct a stack of super-resolution images with z-axis sectioned every 10 nm using QuickPALM14. Adjust the brightness of the stacks and duplicate the stack of one target cell. Use a plugin called 3D Viewer (Table of Materials)15 in ImageJ to generate the 3-D image of the cell. Record the rotation of the 3-D structure for demonstration purposes.</li>
</ol>

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The authors have nothing to disclose.

In this protocol, we described a procedure to significantly reduce the autofluorescence of the cyanobacterium Prochlorococcus (Figure 3C) and, then, immunostain the proteins in the cells, which enabled us to utilize STORM to study the 3-D FtsZ ring morphologies in Prochlorococcus (Figure 4). This protocol might be adopted for super-resolution imaging in other photosynthetic organisms.
Previous studies on photosynthet...

Super-resolution microscopies can break the diffraction limit of light and provide images within sub-diffraction resolutions (&#60; 200 nm). They have been widely used in many organisms to study protein localization and subcellular structures. Major super-resolution microscopy methods include structured illumination microscopy (SIM), stimulated emission depletion microscopy (STED), STORM, and photoactivated localization microscopy (PALM). The mechanisms and applications of these super-resolution microscopes have been reviewed elsewhere1,2.
STORM can achieve a resolution as high as 10 nm by spatial separation3,4. For STORM, only one molecule within a diffraction-limited region is activated (&#34;on&#34;) and the rest of the molecules are kept inactivated (&#34;off&#34;). By an accumulation of rapid switch-on and -off of single molecules, a &#34;diffraction-unlimited&#34; image can be generated3. Meanwhile, many kinds of organic dyes and fluorescent proteins are applicable in STORM, allowing an easy upgrade from regular fluorescence microscopy to high-resolution microscopy5,6.
STORM has not been widely applied in photosynthetic cells, such as cyanobacteria, algae, and plant cells with chloroplasts7,8, which is due to the fact that STORM requires high laser intensity to drive photoswitching. The high-intensity laser unfavorably excites strong autofluorescence background in photosynthetic cells and interferes with the single-molecule localization in STORM imaging. In order to use STORM to investigate the subcellular structures or protein interactions in photosynthetic cells, we developed a photobleaching protocol to quench the background autofluorescence signals9. In a routine immunofluorescent staining procedure, specimens are exposed to white light of a high intensity during the blocking step, which lowers the autofluorescence of photosynthetic cells to meet the requirements for STORM. Thus, this protocol makes it feasible to investigate pigmented organisms with STORM.
Here, we describe the protocol to use STORM to image the FtsZ ring organization in the unicellular picocyanobacterium Prochlorococcus. FtsZ is a highly conserved tubulin-like cytoskeletal protein which polymerizes to form a ring structure (the Z ring) around the circumference of a cell10 and is essential for the cell division11. Preserved Prochlorococcus cells are first photobleached to reduce the autofluorescent background and immunostained with a primary anti-FtsZ antibody, and then a secondary anti-Rabbit IgG (H+L) antibody is conjugated with a fluorophore (e.g., Alexa Fluor 750). Eventually, STORM is used to observe the detailed FtsZ ring organizations in Prochlorococcus during different cell cycle stages.

<table><tbody><tr><td>Polystyrene particles</td><td>Spherotech</td><td>PP-20-10</td><td>2.0-2.4 &amp;micro;m</td></tr><tr><td>Coverslip</td><td>Marienfeld</td><td>0111580</td><td>18 mm &amp;empty;, Thickness No. 1</td></tr><tr><td>Ethanol</td><td>Scharlau</td><td>ET00021000</td><td></td></tr><tr><td>Poly-L-lysine hydrobromide</td><td>Sigma-Aldrich</td><td>P9155</td><td>mol wt 70,000-150,000</td></tr><tr><td>Paraformaldehyde</td><td>Sigma-Aldrich</td><td>158127</td><td></td></tr><tr><td>Glutaraldehyde solution, 50%</td><td>Sigma-Aldrich</td><td>340855</td><td></td></tr><tr><td>PBS</td><td>Sigma</td><td>P3813</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma</td><td>T8787</td><td></td></tr><tr><td>EDTA Disodium Salt, 2-hydrate</td><td>Gold biotechnology</td><td>E-210-500</td><td></td></tr><tr><td>Trizma base</td><td>Sigma</td><td>T1503</td><td></td></tr><tr><td>Lysozyme</td><td>Sigma</td><td>L6876</td><td></td></tr><tr><td>Goat serum</td><td>Sigma</td><td>G9023</td><td></td></tr><tr><td>anti-&lt;em&gt;Anabaena&lt;/em&gt; FtsZ antibody</td><td>Agrisera</td><td>AS07217</td><td></td></tr><tr><td>Goat anti-Rabbit IgG (H+L) Cross-Adsorbed Secondary Antibody</td><td>Life Technologies</td><td>A-21039</td><td>conjugated with Alexa Fluor 750</td></tr><tr><td>D-Glucose Anhydrous</td><td>Fisher Scientific</td><td>D16-1</td><td></td></tr><tr><td>L-Ascorbic Acid</td><td>Sigma-Aldrich</td><td>A5960</td><td></td></tr><tr><td>Methyl Viologen</td><td>Sigma-Aldrich</td><td>856177</td><td></td></tr><tr><td>Cyclooctatetraene</td><td>Sigma-Aldrich</td><td>138924</td><td></td></tr><tr><td>tris(2-carboxyethyl)phosphine (TCEP)</td><td>Sigma-Aldrich</td><td>646547</td><td></td></tr><tr><td>Glucose Oxidase</td><td>Sigma-Aldrich</td><td>G2133</td><td></td></tr><tr><td>Catalase</td><td>Sigma-Aldrich</td><td>C9322</td><td></td></tr><tr><td>XD-300 Xenon light source</td><td></td><td></td><td>250 W</td></tr><tr><td>STORM microscope</td><td>NBI</td><td>SRiS microscope</td><td></td></tr><tr><td>Rohdea</td><td>NBI</td><td>SRiS 3.0</td><td>software for imaging acquisition</td></tr><tr><td>Luna</td><td>NBI</td><td>SRiS 3.0</td><td>software for drifting correction</td></tr><tr><td>QuickPALM</td><td></td><td></td><td>https://code.google.com/archive/p/quickpalm/wikis</td></tr><tr><td>3D Viewer</td><td></td><td></td><td>http://132.187.25.13/ij3d/?page=Home&amp;amp;category=Home</td></tr></tbody></table>

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.

STORM achieves super-resolution imaging by activating individual photoswitchable fluorophores stochastically. The location of every fluorophore is recorded and a super-resolution image is then constructed based on these locations4. Therefore, the precision of the fluorophore location is important for the super-resolution image reconstruction. The absorption spectra of Prochlorococcus peak at 447 nm and 680 nm,and Prochlorococcus has a minimum abso...

Watch this Scientific Journal Video about Photobleaching Enables Super-resolution Imaging of the FtsZ Ring in the Cyanobacterium Prochlorococcus at JoVE.com

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.

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, ...

photobleaching-enables-super-resolution-imaging-ftsz-ring

Research

JoVE Journal

Immunology and Infection

6.2K Views.  Hong Kong University of Science and Technology. Deze methode kan helpen bij het beantwoorden van belangrijke vragen over eiwitlokalisatie en de dynamiek in fotosynthetische organismen. Door de fotobleachingmethode te combineren met een microscoop met superresolutie, zijn we in staat om de eiwitdynamiek van fotosynthetische organismen tot in de kleinste details te detecteren. Deze methode geeft niet alleen inzicht in de eiwitdynamiek van Prochlorococcus.Het kan ook worden toegepast op vele andere organismen die radio-re-dop-sing en b...

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

Multimodal Optical Microscopy Methods Reveal Polyp Tissue Morphology and Structure in Caribbean Reef Building Corals

An integrated suite of imaging techniques has been applied to determine the three-dimensional (3D) morphology and cellular structure of polyp tissues comprising the Caribbean reef building corals Montastraeaannularis and M. faveolata. These approaches include fluorescence microscopy (FM), serial block face imaging (SBFI), and two-photon confocal laser scanning microscopy (TPLSM). SBFI provides deep tissue imaging after physical sectioning; it details the tissue surface texture and 3D visualization to tissue depths of more than 2 mm. Complementary FM and TPLSM yield ultra-high resolution images of tissue cellular structure. Results have: (1) identified previously unreported lobate tissue morphologies on the outer wall of individual coral polyps&#160;and (2) created the first surface maps of the 3D distribution and tissue density of chromatophores and algae-like dinoflagellate zooxanthellae endosymbionts. Spectral absorption peaks of 500 nm and 675 nm, respectively, suggest that M. annularis and M. faveolata contain similar types of chlorophyll and chromatophores. However, M. annularis and M. faveolata exhibit significant differences in the tissue density and 3D distribution of these key cellular components. This study focusing on imaging methods indicates that SBFI is extremely useful for analysis of large mm-scale samples of decalcified coral tissues. Complimentary FM and TPLSM reveal subtle submillimeter&#160;scale changes in cellular distribution and density in nondecalcified coral tissue samples. The TPLSM technique affords: (1) minimally invasive sample preparation,&#160;(2) superior optical sectioning ability,&#160;and (3) minimal light absorption and scattering, while still permitting deep tissue imaging.

An integrated suite of imaging techniques has been applied to determine polyp morphology and tissue structure in the Caribbean corals Montastraeaannularis and M. faveolata. Fluorescence, serial block face,&#160;and two-photon confocal laser scanning microscopy have identified lobate structure, polyp walls, and estimated chromatophore and zooxanthellae densities and distributions.

An integrated suite of imaging techniques has been applied to determine the three-dimensional (3D) morphology and cellular structure of polyp tissues ...

Environment

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Developmental Biology

Visualization of Cortex Organization and Dynamics in Microorganisms, using Total Internal Reflection Fluorescence Microscopy

TIRF microscopy has emerged as a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules in vitro and in living cells1. The optical phenomenon of total internal reflection occurs when light passes from a medium with high refractive index into a medium with low refractive index at an angle larger than a characteristic critical angle (i.e. closer to being parallel with the boundary). Although all light is reflected back under such conditions, an evanescent wave is created that propagates across and along the boundary, which decays exponentially with distance, and only penetrates sample areas that are 100-200 nm near the interface2. In addition to providing superior axial resolution, the reduced excitation of out of focus fluorophores creates a very high signal to noise ratios and minimizes damaging effects of photobleaching2,3. Being a widefield technique, TIRF also allows faster image acquisition than most scanning based confocal setups.
At first glance, the low penetration depth of TIRF seems to be incompatible with imaging of bacterial and fungal cells, which are often surrounded by thick cell walls. On the contrary, we have found that the cell walls of yeast and bacterial cells actually improve the usability of TIRF and increase the range of observable structures4-8. Many cellular processes can therefore be directly accessed by TIRF in small, single-cell microorganisms, which often offer powerful genetic manipulation techniques. This allows us to perform in vivo biochemistry experiments, where kinetics of protein interactions and activities can be directly assessed in living cells.
We describe here the individual steps required to obtain high quality TIRF images for Saccharomyces cerevisiae or Bacillus subtilis cells. We point out various problems that can affect TIRF visualization of fluorescent probes in cells and illustrate the procedure with several application examples. Finally, we demonstrate how TIRF images can be further improved using established image restoration techniques.

Total Internal Reflection Fluorescence (TIRF) microscopy is a powerful approach to observe structures close to the cell surface at high contrast and temporal resolution. We demonstrate how TIRF can be employed to study protein dynamics at the cortex of cell wall-enclosed bacterial and fungal cells.

TIRF microscopy has emerged as  a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules  in vitro and in living cells1. ...

Bioengineering

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 ...

Biochemistry

Super-resolution Imaging of the Bacterial Division Machinery

Bacterial cell division requires the coordinated assembly of more than ten essential proteins at midcell1,2. Central to this process is the formation of a ring-like suprastructure (Z-ring) by the FtsZ protein at the division plan3,4. The Z-ring consists of multiple single-stranded FtsZ protofilaments, and understanding the arrangement of the protofilaments inside the Z-ring will provide insight into the mechanism of Z-ring assembly and its function as a force generator5,6. This information has remained elusive due to current limitations in conventional fluorescence microscopy and electron microscopy. Conventional fluorescence microscopy is unable to provide a high-resolution image of the Z-ring due to the diffraction limit of light (~200 nm). Electron cryotomographic imaging has detected scattered FtsZ protofilaments in small C. crescentus cells7, but is difficult to apply to larger cells such as E. coli or B. subtilis. Here we describe the application of a super-resolution fluorescence microscopy method, Photoactivated Localization Microscopy (PALM), to quantitatively characterize the structural organization of the E. coli Z-ring8.
	PALM imaging offers both high spatial resolution (~35 nm) and specific labeling to enable unambiguous identification of target proteins. We labeled FtsZ with the photoactivatable fluorescent protein mEos2, which switches from green fluorescence (excitation = 488 nm) to red fluorescence (excitation = 561 nm) upon activation at 405 nm9. During a PALM experiment, single FtsZ-mEos2 molecules are stochastically activated and the corresponding centroid positions of the single molecules are determined with &lt;20 nm precision. A super-resolution image of the Z-ring is then reconstructed by superimposing the centroid positions of all detected FtsZ-mEos2 molecules.
	Using this method, we found that the Z-ring has a fixed width of ~100 nm and is composed of a loose bundle of FtsZ protofilaments that overlap with each other in three dimensions. These data provide a springboard for further investigations of the cell cycle dependent changes of the Z-ring10 and can be applied to other proteins of interest.

We describe a super-resolution imaging method to probe the structural organization of the bacterial FtsZ-ring, an essential apparatus for cell division. This method is based on quantitative analyses of photoactivated localization microscopy (PALM) images and can be applied to other bacterial cytoskeletal proteins.

Bacterial cell division requires the coordinated  assembly of more than ten essential proteins at midcell1,2. Central  to this process is the formation of ...

Biology

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy (f3D-SIM)

Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the diffraction of light allowing super-resolution of live samples. There are currently three main types of super-resolution techniques &#8211; stimulated emission depletion (STED), single-molecule localization microscopy (including techniques such as PALM, STORM, and GDSIM), and structured illumination microscopy (SIM). While STED and single-molecule localization techniques show the largest increases in resolution, they have been slower to offer increased speeds of image acquisition. Three-dimensional SIM (3D-SIM) is a wide-field fluorescence microscopy technique that offers a number of advantages over both single-molecule localization and STED. Resolution is improved, with typical lateral and axial resolutions of 110 and 280 nm, respectively and depth of sampling of up to 30 &#181;m from the coverslip, allowing for imaging of whole cells. Recent advancements (fast 3D-SIM) in the technology increasing the capture rate of raw images allows for fast capture of biological processes occurring in seconds, while significantly reducing photo-toxicity and photobleaching. Here we describe the use of one such method to image bacterial cells harboring the fluorescently-labelled cytokinetic FtsZ protein to show how cells are analyzed and the type of unique information that this technique can provide.

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy f3D-SIM

Spatiotemporal information about dynamic proteins inside live cells is crucial for understanding biology. A type of super-resolution microscopy called fast 3D-structured illumination microscopy (f3D-SIM) reveals unique information about the cytokinetic Z ring in bacteria: both its bead-like appearance and the rapid dynamics of FtsZ within the ring.

Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the ...

Isolation and Fluorescence Imaging for Single-particle Reconstruction of Chlamydomonas Centrioles

Centrioles are large macromolecular assemblies important for the proper execution of fundamental cell biological processes such as cell division, cell motility, or cell signaling. The green algae Chlamydomonas reinhardtii has proven to be an insightful model in the study of centriole architecture, function, and protein composition. Despite great advances toward understanding centriolar architecture, one of the current challenges is to determine the precise localization of centriolar components within structural regions of the centriole in order to better understand their role in centriole biogenesis. A major limitation lies in the resolution of fluorescence microscopy, which complicates the interpretation of protein localization in this organelle with dimensions close to the diffraction limit. To tackle this question, we are providing a method to purify and image a large number of C. reinhardtii centrioles with different orientations using super-resolution microscopy. This technique allows further processing of data through fluorescent single-particle averaging (Fluo-SPA) owing to the large number of centrioles acquired. Fluo-SPA generates averages of stained C. reinhardtii centrioles in different orientations, thus facilitating the localization of distinct proteins in centriolar sub-regions. Importantly, this method can be applied to image centrioles from other species or other large macromolecular assemblies.

Isolation and Fluorescence Imaging for Single-particle Reconstruction of Chlamydomonas Centrioles

We have developed a strategy to purify and image a large number of centrioles in different orientations amenable for super-resolution microscopy and single-particle averaging.

Centrioles are large macromolecular assemblies important for the proper execution of fundamental cell biological processes such as cell division, cell ...

Isolation and Characterization of Intact Phycobilisome in Cyanobacteria

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

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

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

Autofluorescence Imaging to Evaluate Red Algae Physiology

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

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

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

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

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

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

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

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

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