This research was carried out as part of Projects TIN2015-68454-R and 20961/PI/18, financed by the Spanish Ministry of Economy and Competitiveness and the S&#233;neca Foundation of the Murcia Region. Irene Hern&#225;ndez Mart&#237;nez and Francisco Javier Ib&#225;&#241;ez L&#243;pez from the Statistical Support Section of the Scientific and Research Area of Murcia University (Secci&#243;n de Apoyo Estad&#237;stico (SAE), &#193;rea Cient&#237;fica y de Investigaci&#243;n (ACTI), Universidad de Murcia, (Figure 1 were drawn by using pictures from Servier Medical Art. Servier Medical Art by Servier is licensed with a Creative Commons Attribution 3.0 Unported License (https://creativecommons.org/licenses/by/3.0/)

The algal species Chroothece mobilis was used for the present study. The species was obtained from the Microalgae Edaphic SE Spain, MAESE 20.29 culture collection. An overview of the protocol is shown in Figure 1.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64533/64533fig01.jpg" /> 
Figure 1: Overview of the study. Chroothece mobilis is incubated under extreme habitat conditions, such as different monochromatic lights, for 2 weeks. The effect on Chroothece&#39;s physiology is evaluated by autofluorescence of the proteins contained in phycobilisome and photosystems using a confocal laser scanning microscope. <a href="https://www.jove.com/files/ftp_upload/64533/64533fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
1. Sample preparation
<ol>
	<li>Prepare the inoculum of Chroothece mobilis from the agar culture of the collection by transferring it to the SWES liquid medium (Table 2). 
	NOTE: SWES &#34;Seewasser + Erddekokt + Salze&#34; = seawater medium23.</li>
	<li>Maintain all the cultures for 2 weeks with a 16:8 light/dark photoperiod at 20 &#176;C under low white light intensity (LL: 80 &#181;M/m2/s) conditions and without shaking, until the desired cell density is obtained (see step 1.3). 
	NOTE: The growth light conditions are 80 &#181;M/m2/s light intensity (active photosynthetic radiation, PAR). These conditions are used as the low light conditions (control). The composition of the SWES medium is provided in Table 2.</li>
	<li>Carry out inoculations for the different experiments in the exponential culture phase with a cell density of 5 x 103 cells/mL in a 24-well plate, using 1 mL per well. 
	NOTE: Perform cell counting with a Neubauer chamber24, and dilute the culture with SWES whenever necessary.</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2">SWES medium composition</td>
		</tr>
		<tr>
			<td>Component</td>
			<td>Concentration</td>
		</tr>
		<tr>
			<td>KNO3</td>
			<td>1.98 mM</td>
		</tr>
		<tr>
			<td>K2HPO4</td>
			<td>115 &#181;M</td>
		</tr>
		<tr>
			<td>MgSO4</td>
			<td>81 &#181;M</td>
		</tr>
		<tr>
			<td>ZnSO4, 7H2O</td>
			<td>17 nM</td>
		</tr>
		<tr>
			<td>MnSO4, 7H2O</td>
			<td>45 Nm</td>
		</tr>
		<tr>
			<td>H3BO3, 4H2O</td>
			<td>3.1 mM</td>
		</tr>
		<tr>
			<td>Co(NO3)2</td>
			<td>17 mM</td>
		</tr>
		<tr>
			<td>Na2MoO4, 6H2O</td>
			<td>21 nM</td>
		</tr>
		<tr>
			<td>CuSO4, 2H2O</td>
			<td>0.1 nM</td>
		</tr>
		<tr>
			<td>FeSO4, 5H2O</td>
			<td>13 &#181;M</td>
		</tr>
		<tr>
			<td>EDTA, 7H2O</td>
			<td>11 &#181;M</td>
		</tr>
		<tr>
			<td>Vit B12</td>
			<td>5 &#181;g</td>
		</tr>
		<tr>
			<td>Soil extract</td>
			<td>30 mL</td>
		</tr>
		<tr>
			<td>Filtered river water</td>
			<td>455 mL</td>
		</tr>
	</tbody>
</table>
Table 2: SWES medium composition.
2. Reproducing extreme algae habitat conditions: green and red monochromatic light effect
<ol>
	<li>Add 1 mL of the cell culture from the stock culture prepared at the aforementioned cell density (step 1.2) to inoculate each well of a 24-well plate.</li>
	<li>Keep the cell culture covered for 2 weeks to reproduce the monochromatic light effect.
	<ol>
		<li>Use the green filter that allows green light to pass through from 470 to 570 nm with a peak at the 506 nm wavelength (according to the manufacturer&#39;s specifications; see Table of Materials) and expose it to the culture25.</li>
		<li>Use the red filter that allows red light between 590 and 720 nm and peaks at 678 nm (according to the manufacturer&#39;s specifications; see Table of Materials) to expose it to the culture25. 
		NOTE: The low white light intensity (LL: 80 &#181;M/m2/s; see Table of Materials) condition is used as the light intensity control to compare the obtained effects. The employed light intensity units are micromole per second and square meter (&#181;mol m-2 s-1) or photosynthetic photon flux density (PPFD).</li>
	</ol>
	</li>
</ol>
3. Autofluorescence imaging
NOTE: The setup of the imaging software (see Table of Materials) is illustrated in Figure 2.
<ol>
	<li>Turn on all the components of the inverted confocal laser scanning microscope (CLSM; see Table of materials), including the laser.</li>
	<li>Mount the cells from each experimental well of the 24-well plate in SWES growth medium to a 35 mm glass bottom dish (see Table of Materials) for imaging.</li>
	<li>Choose the 63x/1.30 NA glycerol immersion objective and place glycerol over the lens (see Table of Materials).</li>
	<li>Place the well plate on the microscope stage and ensure the specimen does not move during image acquisition.</li>
	<li>Center the specimen in the light path and focus on the center of the cell by selecting the plane with the highest fluorescence intensity.</li>
	<li>Open the image acquisition software and choose xy&#955; from the drop-down list in the acquisition mode26.</li>
	<li>Select the excitation line of the laser to 561 nm DPSS, 8 bits of dynamic range, and 1024 x 1024 pixels. 
	NOTE: Ensure the confocal microscope has a 561 nm DPSS laser or a white light laser (WLL).</li>
	<li>Collect the fluorescence emission spectra in the 10 nm bandwidth and the lambda step size of 4 nm within the 570-760 nm range.</li>
	<li>Set the pinhole at 1 Airy unit and run the lambda scan acquisition.</li>
	<li>Repeat this process as many times as possible in different fields of view to collect an acceptable amount of data for statistical analysis (usually a standard deviation lower than 10%22).</li>
	<li>Repeat the last step under the different conditions (under red and green lights) and save the data. 
	NOTE: Use the same acquisition settings for the different samples and conditions to compare and conduct the statistical analysis.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64533/64533fig02.jpg" /> 
Figure 2: Software setup. Imaging software user interface to set up the lambda scan parameters. (A) From left to right, to select the acquisition mode xy&#955; from the drop-down list, that corresponds to step 3.6 in the protocol, and to select the right immersion lens type, corresponding to step 3.3 in the protocol. Ensure to remove any filter from the light path in step 3.9. (B) The panel for setting up the lambda scan parameters corresponds to step 3.8 in the protocol. (C) Run the lambda scan in step 3.10. <a href="https://www.jove.com/files/ftp_upload/64533/64533fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Parameters to evaluate Chroothece&#39;s physiology
<ol>
	<li>Once the lambda scan is acquired, click on the quantify window at the top of the software (as shown in Figure 3) to evaluate the collected fluorescence emission spectra. Go to the Open project window and select one xy&#955; file (Figure 3).</li>
	<li>Select Stack profile analysis in the imaging software.</li>
	<li>Define a region of interest (ROI) of 4 &#181;m2 in the center of a cell to analyze the mean fluorescence intensity (MFI). Export the data in CSV format. 
	NOTE: It is important to avoid the presence of black pixels (to ensure the pixels selected contain positive values) and always maintain a same sized ROI.</li>
	<li>Repeat this process with different cells under different conditions to produce enough data to perform statistical analysis.</li>
	<li>Open the CSV files to select the different fluorescence emission peaks from the phycobiliproteins and chlorophylls of all the measured ROIs.
	<ol>
		<li>Select the fluorescence data of phycoerythrin-phycocyanobilin (PE-PCB; 620 nm), C-phycocyanin (CPC; 648 nm), allophycocyanin (APC; 660 nm), and chlorophyll a (Chl a; 680 nm), respectively, in the csv file.</li>
	</ol>
	</li>
	<li>Create a new table with all the maximum fluorescence values obtained from each phycobiliprotein and chlorophyll peak and plot the data on a graph.</li>
	<li>Perform statistical analysis.
	<ol>
		<li>Analyze if the obtained data meets normality and homoscedasticity27.</li>
		<li>Proceed to the t-test analysis27,28 if the first condition is true. If not, run a Mann-Whitney U test29.</li>
		<li>Consider the differences significant at p values &#60;0.05.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64533/64533fig03.jpg" /> 
Figure 3: Evaluating Chroothece&#39;s autofluorescence. To perform the autofluorescence analysis, ensure to select the quantify window and one xy&#955; file in the open projects window (step 4.1); select stack profile visualization (step 4.2); select a 4 &#181;m2 ROI in the center of a cell and click on the report button to export the lambda scan data in CSV format (step 4.3). <a href="https://www.jove.com/files/ftp_upload/64533/64533fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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The authors have no conflicts of interest to declare.

Some unicellular or colonial red algae, such as Chroothece,&#160;grow slowly in vitro, but contain multiple autofluorescent compounds that can be analyzed by spectral analysis under a confocal microscope, where differences in pigment emission peaks can be detected. Spectral confocal fluorescence microscopy has allowed us to conduct in vivo studies to evaluate the adaptation or acclimation of photosynthetic organisms8,10,<sup ...

Red algae, such as the genus Chroothece, can grow in extreme habitats, where they frequently have to cope with marked environmental changes1. Floods and droughts are frequent in semi-arid regions where this genus can be found, and some species have been reported in creeks, cliffs, caves, or even thermal waters2. However, most of the time, biological variables, such as competition or grazing, relegate species to non-optimal conditions for their growth. As these organisms are often difficult to culture and either do not grow or grow very slowly under laboratory conditions, one major limitation is the available sample size. Therefore, it is very important to follow non-destructive methods or methods that involve minimal sample manipulation3,4.
The physiological skills needed to survive in these harsh environments can be monitored by following changes in their photosynthetic systems. Metabolic mechanisms, photosynthetic efficiency, and sensitivity to light or culture conditions can be revealed by pigment fluorescence emission profiles, due to accurate changes in their energy transfer or trapping5,6,7,8.
Autofluorescence of cellular compounds can be used as a marker for cytodiagnosis or as a natural indicator of cellular state or metabolism in response to external and internal signals through changes in emission9. It can also be used to discriminate taxonomically different groups of photosynthetic organisms10. Depending on the phylogenetic position of phototrophic microorganisms, one can find different in vivo fluorescence features. Therefore, a taxonomic identification based on the in vivo characteristics of phototrophic fluorescence (including fluorescence absorption and emission spectra) has been attempted on several occasions11,12. Because of the diversity in accessory pigments among phytoplankton taxa, differences in the wavelengths at which chlorophyll a (Chl a) fluorescence is stimulated, or differences in emission spectra, can be used to infer taxonomy13. The in vivo fluorescence excitation and emission spectra of these specimens rely not only on the phyla of algae, but also on photosystem adaptation14. The efficiency of energy transfer to Chl a, or the ratio of Chl a to accessory pigments, and the cellular pigment content are sensitive to growth conditions5.
Red algae, particularly Chroothece, have several accessory fluorescent pigments-phycobiliproteins and carotenoids; the former concentrate in phycobilisomes attached to the thylakoids of chloroplasts. Phycobiliproteins (phycocyanin, phycoerythrin, and allophycocyanin) can capture light at different wavelengths and transmit it to Chl a, which makes photosynthesis in very different light and culture conditions possible15. For instance, Chroothece species can grow inside caves or almost emerge in slightly saline calcareous streams2.
Monochromatic lights affect the growth and pigment composition of photosynthetic organisms, and have been studied to prevent or control the growth of photosynthetic organisms in caves. Mulec et al. showed that red enriched lighting promotes the growth of cyanobacteria, algae, and plants16. Previous studies have also reported that green light affects the pigment composition of cyanobacteria17, while others have revealed that green light prevents the growth of most photosynthetic organisms and some cyanobacteria exhibit a reduction in thylakoids and weaker mean fluorescence intensity18.
To understand the ability of Chroothece as a model organism to overcome harsh conditions, cultured cells have been exposed to increasing light intensities and monochromatic light (green or red)15, to see how it copes with the dim conditions of caves (where red light predominates). The protocol presented herein reproduces the effect of the abovementioned variables on Chroothece&#39;s phycobiliproteins at the cellular level using its own autofluorescence.
Nowadays, fluorescence is commonly used as a tool to study the physiological responses of vascular plants, microalgae, macroalgae, and cyanobacteria13,14,16. Spectral confocal fluorescence microscopy is a superb tool for in vivo studies to evaluate photosynthetic specimens&#39; physiology at the single-cell level10,17,18,19,20, by avoiding problems associated with the low growth rate in the laboratory and the difficulties with obtaining enough biomass for the associated extraction and biochemical methods8. Once cells are treated under different culture conditions for 2 weeks, the lambda scan profile can be measured in vivo. Although there are several publications in which different wavelengths of excitation by confocal imaging have been used3,4,10,17, most phycobiliproteins and Chl a can be detected using a 561 nm wavelength excitation line, and the detected emission ranges from 570 to 760 nm wavelength. These criteria have been based on an analysis previously performed10 with commercial pure pigments (Table 1) by confocal imaging and the obtained results in different algae species20,21,22.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td rowspan="2">Pigments</td>
			<td rowspan="2">&#955;flmax (nm)</td>
			<td colspan="8">&#955; exc (nm)</td>
		</tr>
		<tr>
			<td>351</td>
			<td>364</td>
			<td>458</td>
			<td>476</td>
			<td>488</td>
			<td>514</td>
			<td>543</td>
			<td>633</td>
		</tr>
		<tr>
			<td>Chl a</td>
			<td>660.9-678.1</td>
			<td>43.4 &#177; 1.8</td>
			<td>11.2 &#177; 0.2</td>
			<td>1.8 &#177; 0.05</td>
			<td>2.0 &#177; 0.08</td>
			<td>12.2 &#177; 0.7</td>
			<td>6.0 &#177; 0.3</td>
			<td>4.2 &#177; 0.16</td>
			<td>80.7 &#177; 1.5</td>
		</tr>
		<tr>
			<td rowspan="2">R-PE</td>
			<td>569.2-583.3</td>
			<td>5.9 &#177; 0.6</td>
			<td>5.9 &#177; 0.16</td>
			<td>11.1 &#177; 0.04</td>
			<td>42.2 &#177; 0.3</td>
			<td>100.0 &#177; 0</td>
			<td>90.0 &#177; 0.3</td>
			<td>99.2 &#177; 0.08</td>
			<td>-</td>
		</tr>
		<tr>
			<td>652.1-668.6</td>
			<td>-</td>
			<td>-</td>
			<td>1.5 &#177; 0.01</td>
			<td>3.7 &#177; 0.04</td>
			<td>26.7 &#177; 0.5</td>
			<td>8.7 &#177; 0.16</td>
			<td>11.1 &#177; 0.16</td>
			<td>11.3 &#177; 0.2</td>
		</tr>
		<tr>
			<td>C-PC</td>
			<td>636.2-676.4</td>
			<td>2.3 &#177; 0.04</td>
			<td>1.0 &#177; 0.01</td>
			<td>0.6 &#177; 0.004</td>
			<td>0.7 &#177; 0.008</td>
			<td>2.0 &#177; 0.08</td>
			<td>2.0 &#177; 0.04</td>
			<td>3.3 &#177; 0.16</td>
			<td>33.6 &#177; 0.9</td>
		</tr>
		<tr>
			<td>APC-XL</td>
			<td>667.3-683.8</td>
			<td>15.1 &#177; 1.5</td>
			<td>9.6 &#177; 0.98</td>
			<td>1.0 &#177; 0.04</td>
			<td>1.2 &#177; 0.08</td>
			<td>5.9 &#177; 0.7</td>
			<td>4.1 &#177; 0.5</td>
			<td>23.2 &#177; 3.5</td>
			<td>91.4 &#177; 2.3</td>
		</tr>
	</tbody>
</table>
Table 1: The pure pigment information used to run the lambda scan analysis. This table shows emission peaks and shoulders/fluorescence band maxima of different fluorochromes/pigments by confocal imaging spectrophotometry for all the excitation wavelengths, and the percentage of light emission by pigments/fluorochromes. Values were calculated by the formula: = MFI*100/255. Each value is the mean &#177; SE (mean &#177; standard error from the mean). Pure pigments were used for calibrating the confocal scanning laser microscope as follows1,2,10. Chlorophyll a was obtained from Spinacia oleracea, R-phycoerythrin (R-PE) from Porphyra tenera, and C-phycocyanin (C-PE) from Spirulina sp. All the species were dissolved in filtered distilled water. Allophycocyanin-XL (APC-XL) was obtained from Mastigocladus laminosus, which was dissolved in ammonium sulfate (60%) and potassium phosphate (pH = 7) to achieve a concentration of 38 mM. The scans were performed with 400 &#181;L of each pigment solution (concentration of 1 mg/mL) using an 8-well covered-glass bottom chamber.
The study of a single excitation wavelength is quite a useful first approximation. In this case, however, it is necessary to elucidate the relative contribution of the different complexes in the fluorescence signal, which is recommended to perform a fluorescence ratio or spectrum analysis at several wavelengths, among other methods.

<table><tbody><tr><td>&amp;micro;-Dish 35 mm, high Glass Bottom</td><td>Ibidi&amp;nbsp;</td><td>81158</td><td>-</td></tr><tr><td>24 black well plate</td><td>Ibidi</td><td>82406</td><td>&amp;nbsp;flat and clear bottom for high throughput microscopy</td></tr><tr><td>Algae Incubator</td><td>Panasonic</td><td>MLR-352-PE</td><td></td></tr><tr><td>Confocal laser scanning microscope</td><td>Leica Microsystems</td><td>SP8 TCS</td><td>-</td></tr><tr><td>Flask</td><td>Fisher Scientific</td><td>15380591</td><td>Can be purchased in a local convenience store or online stores.</td></tr><tr><td>green filter</td><td>PNTA, LEE filters</td><td>-</td><td>Can be purchased in a local convenience store or online stores.</td></tr><tr><td>HC PL APO 63X/1.30 GLYC CORR CS2</td><td>Leica Microsystems</td><td>506353</td><td>Glycerol immersion lens</td></tr><tr><td>Image acquisition software. LAS X</td><td>Leica Microsystems</td><td>SP8 TCS</td><td>-</td></tr><tr><td>Light source</td><td>Panasonic</td><td>FL40SSENW/37MLR-352-PE</td><td></td></tr><tr><td>Quantum photoradiometer</td><td>DeltaOhm&amp;nbsp;</td><td>DO 9721</td><td>-</td></tr><tr><td>R software</td><td>R Core Team, 2020</td><td>4.0.2.</td><td>-</td></tr><tr><td>red filter</td><td>PNTA, LEE filters</td><td>-</td><td>Can be purchased in a local convenience store or online stores.</td></tr><tr><td>SWES medium</td><td>University of Murcia</td><td>-</td><td>-</td></tr><tr><td>Type G Immersion liquid</td><td>Leica Microsystems</td><td>11513910</td><td>Glycerol&amp;nbsp;</td></tr></tbody></table>

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.

Chlorophyll a generally absorbs blue and red wavelengths of visible light, whereas phycobiliproteins use green, yellow, and orange wavelengths7. The autofluorescence of these pigments makes the first approach to study phycobiliproteins and chlorophyll behavior under experimental and field conditions possible.
By comparing the obtained data and plotting on different graphs, significant changes in mean fluorescence intensity (MFI) can be distinguished (<strong class="xfig...

Watch this Scientific Journal Video about Autofluorescence Imaging to Evaluate Red Algae Physiology at JoVE.com

Autofluorescence Imaging to Evaluate Red Algae Physiology

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 ...

autofluorescence-imaging-to-evaluate-red-algae-physiology

Research

JoVE Journal

Biology

1.3K Views.  University of Murcia. Dit confocale microscopische protocol zal de interpretatie van het ecologische gedrag en de aanpassingen van microalgen uit extreme omgevingen met langzame groei in het laboratorium mogelijk maken met behulp van autofluorescentie van hun pigmenten. Deze techniek is niet invasief en minimaliseert artefacten omdat chemische verbindingen niet worden gebruikt. Het kennen van de prestaties van pigmenten van autotrofen en de bijbehorende groei zal het ontwerpen van de controlemethoden mogelijk make...

Video: Autofluorescence Imaging to Evaluate Red Algae Physiology

A Simple and Rapid Protocol for Measuring Neutral Lipids in Algal Cells Using Fluorescence

Algae are considered excellent candidates for renewable fuel sources due to their natural lipid storage capabilities. Robust monitoring of algal fermentation processes and screening for new oil-rich strains requires a fast and reliable protocol for determination of intracellular lipid content. Current practices rely largely on gravimetric methods to determine oil content, techniques developed decades ago that are time consuming and require large sample volumes. In this paper, Nile Red, a fluorescent dye that has been used to identify the presence of lipid bodies in numerous types of organisms, is incorporated into a simple, fast, and reliable protocol for measuring the neutral lipid content of Auxenochlorella protothecoides, a green alga. The method uses ethanol, a relatively mild solvent, to permeabilize the cell membrane before staining and a 96 well micro-plate to increase sample capacity during fluorescence intensity measurements. It has been designed with the specific application of monitoring bioprocess performance. Previously dried samples or live samples from a growing culture can be used in the assay.

A simple protocol to determine the neutral lipid content of algal cells using a Nile Red staining procedure is described. This time-saving technique offers an alternative to traditional gravimetric-based lipid quantification protocols. It has been designed for the specific application of monitoring bioprocess performance.

Algae are considered excellent candidates for renewable fuel sources due to their natural lipid storage capabilities. Robust monitoring of algal ...

Chemistry

Ratiometric Biosensors that Measure Mitochondrial Redox State and ATP in Living Yeast Cells

Mitochondria have roles in many cellular processes, from energy metabolism and calcium homeostasis to control of cellular lifespan and programmed cell death. These processes affect and are affected by the redox status of and ATP production by mitochondria. Here, we describe the use of two ratiometric, genetically encoded biosensors that can detect mitochondrial redox state and ATP levels at subcellular resolution in living yeast cells. Mitochondrial redox state is measured using redox-sensitive Green Fluorescent Protein (roGFP) that is targeted to the mitochondrial matrix. Mito-roGFP contains cysteines at positions 147 and 204 of GFP, which undergo reversible and environment-dependent oxidation and reduction, which in turn alter the excitation spectrum of the protein. MitGO-ATeam is a F&ouml;rster resonance energy transfer (FRET) probe in which the &epsilon; subunit of the FoF1-ATP synthase is sandwiched between FRET donor and acceptor fluorescent proteins. Binding of ATP to the &epsilon; subunit results in conformation changes in the protein that bring the FRET donor and acceptor in close proximity and allow for fluorescence resonance energy transfer from the donor to acceptor.

We describe the use of two ratiometric, genetically encoded biosensors, which are based on GFP, to monitor mitochondrial redox state and ATP levels at subcellular resolution in living yeast cells.

Mitochondria have roles in many cellular processes, from energy metabolism and calcium homeostasis to control of cellular lifespan and programmed cell ...

Bioengineering

Molecular Probe Optimization to Determine Cell Mortality in a Photosynthetic Organism (Microcystis aeruginosa) Using Flow Cytometry

Microbial subpopulations in field and laboratory studies have been shown to display high heterogeneity in morphological and physiological parameters. Determining the real time state of a microbial cell goes beyond live or dead categories, as microbes can exist in a dormant state, whereby cell division and metabolic activities are reduced. Given the need for detection and quantification of microbes, flow cytometry (FCM) with molecular probes provides a rapid and accurate method to help determine overall population viability. By using SYTOX Green and SYTOX Orange in the model cyanobacteria Microcystis aeruginosa to detect membrane integrity, we develop a transferable method for rapid indication of single cell mortality. The molecular probes used within this journal will be referred to as green or orange nucleic acid probes respectively (although there are other products with similar excitation and emission wavelengths that have a comparable modes of action, we specifically refer to the fore mentioned probes). Protocols using molecular probes vary between species, differing principally in concentration and incubation times. Following this protocol set out on M.aeruginosa the green nucleic acid probe was optimized at concentrations of 0.5 &#181;M after 30 min of incubation and the orange nucleic acid probe at 1 &#181;M after 10 min. In both probes concentrations less than the stated optimal led to an under reporting of cells with membrane damage. Conversely, 5 &#181;M concentrations and higher in both probes exhibited a type of non-specific staining, whereby &#39;live&#39; cells produced a target fluorescence, leading to an over representation of &#39;non-viable&#39; cell numbers. The positive controls (heat-killed) provided testable dead biomass, although the appropriateness of control generation remains subject to debate. By demonstrating a logical sequence of steps for optimizing the green and orange nucleic acid probes we demonstrate how to create a protocol that can be used to analyse cyanobacterial physiological state effectively.

Molecular Probe Optimization to Determine Cell Mortality in a Photosynthetic Organism Microcystis aeruginosa Using Flow Cytometry

Microbial populations contain substantial cell heterogeneity, which can dictate overall behavior. Molecular probe analysis through flow cytometry can determine physiological states of cells, however its application varies between species. This study provides a protocol to accurately determine cell mortality within a cyanobacterium population, without underestimating or recording false positive results.

Microbial subpopulations in field and laboratory studies have been shown to display high heterogeneity in morphological and physiological parameters. ...

Medicine

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a quality control mechanism that can target and selectively degrade cellular material including organelles1-4. For example, damaged or redundant mitochondria (Fig. 1), not disposed of by autophagy, can represent a threat to cellular homeostasis and cell survival. In the yeast, Saccharomyces cerevisiae, nutrient deprivation (e.g., nitrogen starvation) or damage can promote selective turnover of mitochondria by autophagy in a process termed mitophagy 5-9.
We describe a simple fluorescence microscopy approach to assess autophagy. For clarity we restrict our description here to show how the approach can be used to monitor mitophagy in yeast cells. The assay makes use of a fluorescent reporter, Rosella, which is a dual-emission biosensor comprising a relatively pH-stable red fluorescent protein linked to a pH-sensitive green fluorescent protein. The operation of this reporter relies on differences in pH between the vacuole (pH ~ 5.0-5.5) and mitochondria (pH ~ 8.2) in living cells. Under growing conditions, wild type cells exhibit both red and green fluorescence distributed in a manner characteristic of the mitochondria. Fluorescence emission is not associated with the vacuole. When subjected to nitrogen starvation, a condition which induces mitophagy, in addition to red and green fluorescence labeling the mitochondria, cells exhibit the accumulation of red, but not green fluorescence, in the acidic vacuolar lumen representing the delivery of mitochondria to the vacuole. Scoring cells with red, but not green fluorescent vacuoles can be used as a measure of mitophagic activity in cells5,10-12.

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

A robust approach to monitor the delivery of organelles to the acidic lumen of the yeast vacuole for degradation and recycling is described. The method relies on the specific labeling of target organelles with a genetically encoded dual-emission fluorescence pH-biosensor, and visualization of individual cells using fluorescence microscopy.

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a ...

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

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

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

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

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

Immunology and Infection

Autofluorescence Imaging to Evaluate Cellular Metabolism

Cellular metabolism is the process by which cells generate energy, and many diseases, including cancer, are characterized by abnormal metabolism. Reduced nicotinamide adenine (phosphate) dinucleotide (NAD(P)H) and oxidized flavin adenine dinucleotide (FAD) are coenzymes of metabolic reactions. NAD(P)H and FAD exhibit autofluorescence and can be spectrally isolated by excitation and emission wavelengths. Both coenzymes, NAD(P)H and FAD, can exist in either a free or protein-bound configuration, each of which has a distinct fluorescence lifetime-the time for which the fluorophore remains in the excited state. Fluorescence lifetime imaging (FLIM) allows quantification of the fluorescence intensity and lifetimes of NAD(P)H and FAD for label-free analysis of cellular metabolism. Fluorescence intensity and lifetime microscopes can be optimized for imaging NAD(P)H and FAD by selecting the appropriate excitation and emission wavelengths. Metabolic perturbations by cyanide verify autofluorescence imaging protocols to detect metabolic changes within cells. This article will demonstrate the technique of autofluorescence imaging of NAD(P)H and FAD for measuring cellular metabolism.

This protocol describes fluorescence imaging and analysis of the endogenous metabolic coenzymes, reduced nicotinamide adenine (phosphate) dinucleotide (NAD(P)H), and oxidized flavin adenine dinucleotide (FAD). Autofluorescence imaging of NAD(P)H and FAD provides a label-free, nondestructive method to assess cellular metabolism.

Cellular metabolism is the process by which cells generate energy, and many diseases, including cancer, are characterized by abnormal metabolism. Reduced ...

Cellular Redox Profiling Using High-content Microscopy

Reactive oxygen species (ROS) regulate essential cellular processes including gene expression, migration, differentiation and proliferation. However, excessive ROS levels induce a state of oxidative stress, which is accompanied by irreversible oxidative damage to DNA, lipids and proteins. Thus, quantification of ROS provides a direct proxy for cellular health condition. Since mitochondria are among the major cellular sources and targets of ROS, joint analysis of mitochondrial function and ROS production in the same cells is crucial for better understanding the interconnection in pathophysiological conditions. Therefore, a high-content microscopy-based strategy was developed for simultaneous quantification of intracellular ROS levels, mitochondrial membrane potential (&#916;&#936;m) and mitochondrial morphology. It is based on automated widefield fluorescence microscopy and image analysis of living adherent cells, grown in multi-well plates, and stained with the cell-permeable fluorescent reporter molecules CM-H2DCFDA (ROS) and TMRM (&#916;&#936;m and mitochondrial morphology). In contrast with fluorimetry or flow-cytometry, this strategy allows quantification of subcellular parameters at the level of the individual cell with high spatiotemporal resolution, both before and after experimental stimulation. Importantly, the image-based nature of the method allows extracting morphological parameters in addition to signal intensities. The combined feature set is used for explorative and statistical multivariate data analysis to detect differences between subpopulations, cell types and/or treatments. Here, a detailed description of the assay is provided, along with an example experiment that proves its potential for unambiguous discrimination between cellular states after chemical perturbation.

This paper presents a high-content microscopy workflow for simultaneous quantification of intracellular ROS levels, as well as mitochondrial membrane potential and morphology &#8211; jointly referred to as mitochondrial morphofunction &#8211; in living adherent cells using the cell-permeant fluorescent reporter molecules 5-(and-6)-chloromethyl-2&#39;,7&#39;-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H2DCFDA) and tetramethylrhodamine methylester (TMRM).

Reactive oxygen species (ROS) regulate essential cellular processes including gene expression, migration, differentiation and proliferation. However, ...

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

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

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

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

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

Quantifying Retinal Pigment Epithelial Phagocytosis and Lipofuscin Accumulation

Improved Lipofuscin Models and Quantification of Outer Segment Phagocytosis Capacity in Highly Polarized Human Retinal Pigment Epithelial Cultures

The daily phagocytosis of photoreceptor outer segments by the retinal pigment epithelium (RPE) contributes to the accumulation of an intracellular aging pigment termed lipofuscin. The toxicity of lipofuscin is well established in Stargardt's disease, the most common inherited retinal degeneration, but is more controversial in age-related macular degeneration (AMD), the leading cause of irreversible blindness in the developed world. Determining lipofuscin toxicity in humans has been difficult, and animal models of Stargardt's have limited toxicity. Thus, in vitro models that mimic human RPE in vivo are needed to better understand lipofuscin generation, clearance, and toxicity. The majority of cell culture lipofuscin models to date have been in cell lines or have involved feeding RPE a single component of the complex lipofuscin mixture rather than fragments/tips of the entire photoreceptor outer segment, which generates a more complete and physiologic lipofuscin model. Described here is a method to induce the accumulation of lipofuscin-like material (termed undigestible autofluorescence material, or UAM) in highly differentiated primary human pre-natal RPE (hfRPE) and induced pluripotent stem cell (iPSC) derived RPE. UAM accumulated in cultures by repeated feedings of ultraviolet light-treated OS fragments taken up by the RPE via phagocytosis. The key ways that UAM approximates and differs from lipofuscin in vivo are also discussed. Accompanying this model of lipofuscin-like accumulation, imaging methods to distinguish the broad autofluorescence spectrum of UAM granules from concurrent antibody staining are introduced. Finally, to assess the impact of UAM on RPE phagocytosis capacity, a new method for quantifying outer segment fragment/tips uptake and breakdown has been introduced. Termed "Total Consumptive Capacity", this method overcomes potential misinterpretations of RPE phagocytosis capacity inherent in classic outer segment "pulse-chase" assays. The models and techniques introduced here can be used to study lipofuscin generation and clearance pathways and putative toxicity.

Author Spotlight: Improved Lipofuscin Models and Quantification of Outer Segment Phagocytosis Capacity in Highly Polarized Human Retinal Pigment Epithelial Cultures  

This protocol describes a lipofuscin accumulation model in highly differentiated and polarized human retinal pigment epithelial (RPE) cultures and an improved outer segment (OS) phagocytosis assay to detect the total OS consumption/degradation capacity of the RPE. These methods overcome the limitations of previous lipofuscin models and classical pulse-chase outer segment phagocytosis assays.

The daily phagocytosis of photoreceptor outer segments by the retinal pigment epithelium (RPE) contributes to the accumulation of an intracellular aging ...

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 ...

Autofluorescence Imaging to Evaluate Red Algae Physiology

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