J.O.B. and A.M.R. were supported by predoctoral fellowships from the National Science Foundation.

NOTE: For this protocol, we have focused on assaying stromule frequency in the epidermis of N. benthamiana leaves. Several stable transgenic lines have been generated that can be used for this purpose, including 35SPRO:FNRtp:EGFP13 and NRIP1:Cerulean6. Both of these lines show robust expression of fluorophores in the chloroplast stroma of leaves grown under a wide range of conditions. Alternatively, chloroplast-targeted fluorophores may be transiently expressed in N. benthamiana using Agrobacterium transformations13. This is less ideal than the transgenic lines, since Agrobacterium infiltrations induce some basal defense responses in N. benthamiana and interactions with Agrobacterium can alter stromule frequency in the leaf14, potentially complicating interpretation of results. Finally, to visualize stromule formation in vitro, chloroplasts may be extracted from any plant species, using either genetically encoded fluorophores or a fluorescent dye, as described in section 5 below.9,15
NOTE: Detailed methods for plant cultivation have been previously described.16 Briefly, grow N. benthamiana plants in 4&#34; pots filled with any professional soil mix that provides good drainage. Cover seedlings with a clear plastic dome for the first 10-14 days to provide a humid environment for germination. Add any standard fertilizer mix following manufacturer&#39;s instructions to 14-day-old plants. Grow plants under white light, using ~100 &#181;mol photons m-2 sec-1 light intensity. Water plants regularly.
1. Preparing Leaf Samples for Visualization
NOTE: Stromule dynamics are affected by wounding8, so tissue preparation should be conducted immediately before visualizing stromules by confocal fluorescence microscopy. Ideally, a sample should be visualized with 15 min after removal from the plant.
<ol>
	<li>Cut a small section of the leaf using a very sharp razor blade. Always use the same region of the leaf for consistency, and be certain to visualize the same surface (adaxial or abaxial) in all experiments.</li>
	<li>Immediately after cutting the leaf section, submerge the leaf section in a 5 ml needleless syringe filled with water, remove air from the syringe, and apply a vacuum by covering the syringe orifice with a finger and pulling the plunger.
	<ol>
		<li>Release the plunger gently to avoid damaging the leaf section. Repeat this two or three times, or until most of the air has been removed and the leaf appears to be deep green. 
		NOTE: This step removes air in the leaf that can interfere with accurate visualization of stromules.</li>
	</ol>
	</li>
	<li>Add a drop of water to a slide and place the leaf section on this drop. Then, add another drop of water to the top of the leaf, and place a cover slip over the leaf. If there are any air bubbles, gently tap the cover slip until they are removed. The slide is now ready for microscopy, and should be visualized immediately.</li>
</ol>
2. Visualizing Stromules with Confocal Fluorescence Microscopy
<ol>
	<li>With transmitted light, focus on a field of view near the center of the leaf section (away from the cells damaged by the razor blade). Use a 20X objective so that many chloroplasts can be visualized simultaneously. Save an image with transmitted light so that cell types can be distinguished later, if necessary.</li>
	<li>Switch the illumination source to a laser with appropriate excitation and emission filters for the fluorophore being used. For example, excite GFP with a 488 nm laser and collect emissions between 500 nm and 525 nm.
	<ol>
		<li>Using the microscope&#39;s software, select the GFP channel and click to adjust the pinhole aperture to 1 Airy unit. Select laser scanning, to start visualizing the sample, and then click the GFP channel to adjust the laser intensity and the detector gain to minimize the laser power while still clearly visualizing stromules. Once these settings have been determined, keep them as consistent as possible throughout all experiments.</li>
	</ol>
	</li>
	<li>Prepare a z-stack experiment that will collect a series of images through the leaf epidermis in the field of view.
	<ol>
		<li>For the z-stack, include all epidermal chloroplasts in the field of view to avoid bias (for example, if chloroplasts near the leaf surface have more stromules under conditions tested).</li>
		<li>Take images at the maximum interval possible that will include all stromules but minimize the time required for a z-stack. For example, if 1 Airy unit is equivalent to an in-focus confocal section that is 2 &#181;m deep, taking an image every 2 &#181;m is likely ideal. 
		NOTE: The leaf epidermis is an uneven surface, so the total depth of the z-stack will vary depending on the sample; most z-stacks will require 10-20 images, but may include more.</li>
		<li>Adjust scanning speed and image resolution as necessary to make sure that the z-stack is collected quickly (typically within 5-10 min, if possible). Save the z-stack for later analysis.</li>
	</ol>
	</li>
</ol>
3. Image Processing
<ol>
	<li>Determine stromule frequency using any image analysis software. For this protocol, we recommend using ImageJ, which is publicly available from the National Institutes of Health (http://imagej.nih.gov/ij/), or the popular upgrade of ImageJ, Fiji (http://fiji.sc/Fiji).</li>
	<li>Merge the z-stack into a single image using the maximum intensity projection in the software.</li>
	<li>Identify and manually count all chloroplasts in the image.</li>
	<li>For each chloroplast, visually determine whether one or more stromules are seen extending from the chloroplast in the merged z-stack image. For examples, see Figure 1. 
	NOTE: Since the biological functions of stromules are not known, any thin, stroma-filled, tubular extension from the chloroplast can be considered a &#34;stromule&#34;, regardless of length. As a general rule, a stroma-filled extension from the chloroplast is a stromule if it is less than approximately 1 &#181;m in diameter along most of its length7. Ensure that one person analyzes all images to control for subjective differences in determining whether or not a chloroplast has a stromule. A second researcher should independently verify the stromule frequencies to further reduce subjective biases.</li>
</ol>
4. Experimental Design and Sampling
NOTE: Stromule frequency is highly variable between leaves, but several reports suggest that there is little variation in stromule frequency within an individual leaf9,17.
<ol>
	<li>Calculate the leaf stromule frequency from a single field of view of one leaf. Discard the leaf, and consider this an independent sample. Do not consider separate fields of view from one leaf as independent samples. 
	NOTE: There is no strong consensus on how to best measure changes in stromule activity, and until the function of stromules is more clearly defined, researchers should continue to be creative in quantifying stromule dynamics. For comparison across the literature, however, common standards of reporting should be used in addition to more creative approaches. At a minimum, always report the frequency of chloroplasts that have one or more stromules.</li>
	<li>Determine the frequency of chloroplasts that have stromules in a field of view. Use ImageJ to identify all chloroplasts in a field of view. Then, for each chloroplast, determine whether the chloroplast has formed at least one stromule. Report stromule frequency as the percentage of chloroplasts with a stromule. 
	NOTE: Some researchers transform the percentage, for example with the arcsine transformation, before reporting stromule frequencies; while this is an option for statistical analysis, the arcsine of stromule frequency is not an intuitive value, and does not need to be reported in the main text or figures of a study.
	<ol>
		<li>As an alternative approach, count the total number of stromules, and divide it by the total number of chloroplasts. 
		NOTE: Under most circumstances, this will yield similar results to approach 4.2; it may overestimate the stromule frequency, however, if a single chloroplast has formed many stromules. In the example shown under representative results, for example, several chloroplasts have two or more stromules, raising the number of stromules per chloroplast to 40/87 (versus a stromule frequency of 33/87).</li>
		<li>Alternatively, determine the stromule length instead of stromule frequency. Length can be measured in ImageJ by tracing the stromule with the freehand line tool. 
		NOTE: Since the biological role of stromules remains unknown, it is not clear that stromule length affects their function. Report significant differences in stromule length if they are observed, but also report the stromule frequencies (as described in 4.2). 
		NOTE: Stromule frequency can be highly variable among different leaves under the same growth conditions. Therefore, although determining stromule frequency can be time-consuming, experiments should be carefully designed to include large sample sizes. Using datasets from our lab, we determined that a sample size of at least 16 plants for each treatment is usually sufficiently powerful to determine if there is a statistically significant difference of at least a 1.5-fold change in stromule frequency between two conditions (with standard &#945; = 0.05 and &#946; = 0.80).</li>
	</ol>
	</li>
	<li>Statistical analysis of results.
	<ol>
		<li>To compare the mean stromule frequencies of two treatments, use the Welch&#39;s t test (also known as the heteroscedastic t test or t test assuming unequal variance). 
		NOTE: This test is not as powerful as the conventional Student&#39;s t test, but Welch&#39;s t test is advantageous because it does not assume that the variance in stromule frequency distribution is similar between treatments. 
		NOTE: Since stromule frequency is a &#34;proportion&#34;, its sampling distribution is predicted to be binomial rather than normal, which could theoretically skew statistical analysis if sample sizes are very low or if stromule frequency is very low (close to 0%) or very high (close to 100%). Some reports have used arcsine transformations before statistical analysis, which is intended to transform a binomial distribution to a normal distribution, and thus satisfy the standard requirements of a Student&#39;s t test or ANOVA. In practice, however, arcsine transformations have little or no effect on statistical analysis, and are therefore not recommended. Instead, repeat experiments to increase sample size, which will increase statistical power and reduce errors in interpretation of data.</li>
		<li>Whatever statistical approach is used, be sure to carefully report the sampling strategy, sample size, statistical test, and p value for each experiment. 
		NOTE: As with other fields of biology, a p value less than 0.05 may be considered &#34;statistically significant&#34;, although researchers should certainly report the precise p value obtained. If p is slightly above 0.05, increasing the sample size with two or three experiments may help to clarify whether or not the difference observed is reproducible and statistically significant.</li>
	</ol>
	</li>
</ol>
5. Extracting Intact Chloroplasts to Visualize Stromule Dynamics
NOTE: Several methods have been used to isolate chloroplasts from leaves, including a slightly different protocol in a recent study on stromule formation in vitro15. The protocol detailed below uses a relatively simple method that does not yield biochemically pure chloroplast samples, but does instead isolate a large quantity of intact, healthy chloroplasts9,18.
<ol>
	<li>Prepare cold extraction buffer: 50 mM Hepes NaOH, 330 mM sorbitol, 2 mM EDTA, 1.0 mM MgCl2, and 1.0 mM MnCl2. Adjust the pH to 6.9 with NaOH and HCl, and refrigerate before use. 
	NOTE: Although not necessary, using cold buffers and keeping samples on ice when possible will yield a higher proportion of intact chloroplasts.</li>
	<li>Prepare isolation buffer: 50 mM Hepes NaOH, 330 mM sorbitol, 2 mM EDTA, 1.0 mM MnCl2, 1.0 mM MgCl2, 10 mM KCl, and 1.0 mM NaCl. Adjust the pH to 7.6 with NaOH and HCl and refrigerate before use.</li>
	<li>If the leaves are not expressing a genetically encoded stromal fluorophore, prepare carboxyfluorescein diacetate (CFDA) solution: 50 mM CFDA stock solution in dimethyl sulfoxide (DMSO) (2.3 mg CFDA per 1.0 ml DMSO). 
	NOTE: The exact concentration is not critical. Keep CFDA stock in the dark at all times. Store small aliquots of 50 mM CFDA at -20 &#176;C.</li>
	<li>To isolate chloroplasts, remove ~5-10 g leaves from several plants and briefly rinse in cold water. Immediately transfer to 50 ml cold extraction buffer. Grind leaves using a blender with several short pulses. Filter through two or three layers of cheesecloth to remove leaf debris, divide the extracted chloroplasts into two 50 ml centrifuge tubes, and centrifuge for 1 min at 750 x g.</li>
	<li>Discard the supernatant and resuspend the green chloroplasts in 10 ml isolation buffer. Centrifuge again for 1 min at 750 x g, discard supernatant, and resuspend chloroplasts in isolation buffer to bring final volume to 5 ml.</li>
	<li>Transfer 20 &#181;l of the chloroplasts to a slide, cover with a coverslip, and begin microscopy. 
	NOTE: Chloroplasts from transgenic plants expressing plastid-targeted fluorescent proteins are now ready for visualization by confocal fluorescence microscopy.</li>
	<li>Stain chloroplasts from plants that are not expressing plastid-targeted fluorescent proteins to visualize stromules.
	<ol>
		<li>Add 5 &#181;l of 50 mM CFDA stock to the 5 ml chloroplasts-suspended in isolation buffer. Bring the final concentration to 50 &#181;M.</li>
		<li>Allow chloroplasts to incubate for 5 min, and then transfer a small aliquot (~50 &#181;l) to a slide, cover with a coverslip, and begin microscopy.</li>
		<li>Use a FITC or GFP filter set. Use a 20X objective to visualize many chloroplasts simultaneously, or a higher objective to visualize a single isolated chloroplast. 
		NOTE: CFDA will fluoresce inside the stroma of intact chloroplasts.</li>
		<li>If there is strong background fluorescence, wash the chloroplasts again in 10 ml isolation buffer after the 5 min incubation with CFDA, centrifuge for 1 min at 750 x g, discard supernatant, and resuspend in 5 ml isolation buffer.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

When investigating stromules, three important factors must be considered throughout: (i) manipulation of the plant tissue must be kept to an absolute minimum, (ii) the experimental system must be kept consistent, and (iii) sampling strategies must be carefully planned to ensure robust, reproducible data are analyzed.
Stromules are remarkably dynamic: they can extend and retract rapidly before an observer&#39;s eyes under the microscope. Moreover, stromule frequency varies significantly in resp...

Chloroplasts are dynamic organelles in plant cells responsible for photosynthesis and a host of other metabolic processes. Signaling pathways from the chloroplast also exert significant influence on plant physiology and development, coordinating plant responses to environmental stress, pathogens, and even leaf shape1-6. Recently, biologists have gained interest in a poorly understood aspect of chloroplast structure: stromules, very thin stroma-filled tubules that extend from the surface of the chloroplast7.
The biological functions of stromules remain unknown, although stromule frequency is known to vary in response to environmental stimuli7-9, and stromules may be capable of transmitting signaling molecules between organelles6. All types of plastids (not only the green, photosynthetic chloroplasts, but also clear leucoplasts, starch-filled amyloplasts, and pigmented chromoplasts, to name a few types of plastids) make stromules, and stromules are found in all land plant species that have been examined to date. Stromules can extend and retract dynamically, appearing or disappearing within seconds, or they can remain relatively stationary for long times. One of the major hurdles facing stromule biologists is that stromules are often studied using dramatically different methods, tissues, and species, making comparisons across the stromule biology literature difficult. Going forward, standard practices and thorough descriptions of the experimental systems used to study stromules will be critical to discovering the function of these ubiquitous features of chloroplast morphology.
Here we describe methods for visualizing stromule formation in the epidermal chloroplasts of Nicotiana benthamiana leaves. In the mesophyll, chloroplasts are densely packed into large, three-dimensional cells, which makes it difficult to accurately and rapidly visualize stromules by confocal microscopy. By contrast, epidermal cells are relatively flat, contain fewer chloroplasts, and are at the surface of the leaf, allowing for easy and rapid visualization of stromules. N. benthamiana is an ideal model system for these experiments because, unlike many plant species, all cells in the epidermis of N. benthamiana make chloroplasts10. In the epidermis of most plants, including Arabidopsis thaliana, only the stomatal guard cells have chloroplasts, while other epidermal cells have "leucoplasts", plastids that are clear, relatively amorphous, and nonphotosynthetic9,11,12. Thus, whereas a single field of view of an A. thaliana epidermis might show only a handful of chloroplasts in a pair of guard cells, a field of view of an N. benthamiana epidermis will include dozens or even hundreds of chloroplasts. All of the methods described here, however, can be modified to investigate other questions in stromule biology; for example, we have used the same approach to study leucoplast stromules of A. thaliana9.

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	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Hepes</td>
			<td style="width:71px;">Sigma-Aldrich</td>
			<td style="width:119px;">H3375</td>
			<td style="width:167px;"></td>
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			<td height="42" style="height:42px;width:216px;">NaOH</td>
			<td style="width:71px;">Fischer-Scientific</td>
			<td style="width:119px;">S320-1</td>
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			<td height="42" style="height:42px;width:216px;">Sorbitol</td>
			<td style="width:71px;">Sigma-Aldrich</td>
			<td style="width:119px;">S1876</td>
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			<td height="42" style="height:42px;width:216px;">EDTA</td>
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			<td height="42" style="height:42px;width:216px;">MnCl2</td>
			<td style="width:71px;">Sigma-Aldrich</td>
			<td style="width:119px;">221279</td>
			<td></td>
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		<tr height="42">
			<td height="42" style="height:42px;width:216px;">MgCl2</td>
			<td style="width:71px;">Sigma-Aldrich</td>
			<td style="width:119px;">M0250</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">KCl</td>
			<td style="width:71px;">Sigma-Aldrich</td>
			<td style="width:119px;">P3911</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">NaCl</td>
			<td style="width:71px;">Sigma-Aldrich</td>
			<td style="width:119px;">S9625</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Laser Scanning Confocal Microscope&#160;</td>
			<td style="width:71px;">Carl Zeiss Inc</td>
			<td style="width:119px;">Model: LSM710</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Carboxyfluorescein diacetate (CFDA)</td>
			<td style="width:71px;">Sigma-Aldrich</td>
			<td style="width:119px;">21879&#160;</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Dimethyl sulfoxide (DMSO)</td>
			<td style="width:71px;">EMD</td>
			<td style="width:119px;">MX1458-6</td>
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			<td height="21" style="height:21px;width:216px;">Waring blender</td>
			<td style="width:71px;">Waring&#160;</td>
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			<td height="21" style="height:21px;width:216px;">Fiji</td>
			<td style="width:71px;">fiji.sc</td>
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			<td>Open-source software for analyzing biological images</td>
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visualizing stromule frequency with fluorescence microscopy

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but whose functions remain mysterious. Alongside growing attention on the role of chloroplasts in coordinating plant responses to stress, interest in stromules and their relationship to chloroplast signaling dynamics has increased in recent years, aided by advances in fluorescence microscopy and protein fluorophores that allow for rapid, accurate visualization of stromule dynamics. Here, we provide detailed protocols to assay stromule frequency in the epidermal chloroplasts of Nicotiana benthamiana, an excellent model system for investigating chloroplast stromule biology. We also provide methods for visualizing chloroplast stromules in vitro by extracting chloroplasts from leaves. Finally, we outline sampling strategies and statistical approaches to analyze differences in stromule frequencies in response to stimuli, such as environmental stress, chemical treatments, or gene silencing. Researchers can use these protocols as a starting point to develop new methods for innovative experiments to explore how and why chloroplasts make stromules.

This protocol was used to visualize stromule frequency at day and at night in the cotyledons of young N. benthamiana seedlings. Slices from a z-stack were merged into a single image (Figure 1A). For visual purposes, that image was then desaturated and inverted so that stroma appears black (Figure 1B). The chloroplasts were labeled either as having no stromules (green asterisk) or having at least one stromule (magenta asterisk). Of the 87...

Watch this Scientific Journal Video about Visualizing Stromule Frequency with Fluorescence Microscopy at JoVE.com

Visualizing Stromule Frequency with Fluorescence Microscopy

Protocols to investigate the dynamics of chloroplast stromules, the stroma-filled tubules that extend from the surface of chloroplasts, are described.

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but ...

visualizing-stromule-frequency-with-fluorescence-microscopy

Research

JoVE Journal

Biology

8.0K Views.  USDA. Protocols to investigate the dynamics of chloroplast stromules, the stroma-filled tubules that extend from the surface of chloroplasts, are described.

Video: Visualizing Stromule Frequency with Fluorescence Microscopy

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle synapses, we developed a larval tissue preparation that had features of live intact larvae, but also had good optical properties.  This new preparation also allowed for access of perfusates to the synapse.  We used fly larvae, immersed them in artificial hemolymph, and relaxed their normal rhythmic body contractions by chilling them. Next we dissected off the posterior segments of each animal and with a blunt insect pin pushed the mouth parts backward through the body cavity.  This everted the larval body wall, like turning a sock inside-out.  We completed the dissection with ultra-fine dissection scissors and thus exposed the visceral side of the body wall muscles. The glial structures at the NMJ expressed membrane targeted GFP under the control of glial specific promoters. The post-synaptic membrane, the SSR (Subsynaptic Reticula) in muscle expressed synaptically targeted dsRed. We needed to acutely label the motor neuron terminals, the third part of the synapse. To do this we applied primary antibodies to HRP, conjugated to a far-red emitting flurophore.  To test for dye diffusion properties into the perisynaptic space between the motor neuron terminals and the SSR, we applied a solution of large Dextran molecules conjugated to far-red emitting flurophore and collected images.

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

We described structural features of the Glia-neuromuscular synapses in a novel Inside-out tissue preparation of live fly larvae using fluorescent dyes with confocal microscopy. We labeled live neuron terminals with fluorescent primary antibodies to HRP, and also visualized the perisynaptic space with fluorescent Dextrans.

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle ...

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular level. To date the shear complexity of biological systems has caused precise single-molecule experimentation to be far too demanding, instead focusing on studies of single systems using relatively crude bulk ensemble-average measurements. However, many important processes occur in the living cell at the level of just one or a few molecules; ensemble measurements generally mask the stochastic and heterogeneous nature of these events. Here, using advanced optical microscopy and analytical image analysis tools we demonstrate how to monitor proteins within a single living bacterial cell to a precision of single molecules and how we can observe dynamics within molecular complexes in functioning biological machines. The techniques are directly relevant physiologically. They are minimally-perturbative and non-invasive to the biological sample under study and are fully attuned for investigations in living material, features not readily available to other single-molecule approaches of biophysics. In addition, the biological specimens studied all produce fluorescently-tagged protein at levels which are almost identical to the unmodified cell strains ("genomic encoding"), as opposed to the more common but less ideal approach for generating significantly more protein than would occur naturally ('plasmid expression'). Thus, the actual biological samples which will be investigated are significantly closer to the natural organisms, and therefore the observations more relevant to real physiological processes.

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Here we demonstrate the protocols for performing single-molecule fluorescence microscopy on living bacterial cells to enable functional molecular complexes to be detected, tracked and quantified.

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular ...

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Cell Electrofusion Visualized with Fluorescence Microscopy

Cell electrofusion is a safe, non-viral and non-chemical method that can be used for preparing hybrid cells for human therapy. Electrofusion involves application of short high-voltage electric pulses to cells that are in close contact. Application of short, high-voltage electric pulses causes destabilization of cell plasma membranes. Destabilized membranes are more permeable for different molecules and also prone to fusion with any neighboring destabilized membranes. Electrofusion is thus a convenient method to achieve a non-specific fusion of very different cells in vitro. In order to obtain fusion, cell membranes, destabilized by electric field, must be in a close contact to allow merging of their lipid bilayers and consequently their cytoplasm. In this video, we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method. In this method, cells are allowed to attach only slightly to the surface of the well, so that medium can be exchanged and cells still preserve their spherical shape. Fusion visualization is assessed by pre-labeling of the cytoplasm of cells with different fluorescent cell tracker dyes; half of the cells are labeled with orange CMRA and the other half with green CMFDA. Fusion yield is determined as the number of dually fluorescent cells divided with the number of all cells multiplied by two.

In this video we demonstrate efficient electrofusion of cells in vitro by means of modified adherence method using electroporation and the subsequent detection of fused cells visualization with fluorescence microscopy.

Cell electrofusion is a safe, non-viral and non-chemical method that can be used for preparing hybrid cells for human therapy. Electrofusion involves ...

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.

The fission yeast, Schizosaccharomyces pombe, is a good model system to study basic cellular processes. Here we describe a method to perform quantitative live cell analysis of fission yeast. In this particular experiment we focus on organisation of the genome within the cell nucleus, but the method can also be used to study cytosolic factors.

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using ...

Visualizing Bacteria in Nematodes using Fluorescent Microscopy

Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on earth, nematodes and bacteria form a wide array of symbiotic associations that range from beneficial to pathogenic 1-3. One such association is the mutually beneficial relationship between Xenorhabdus bacteria and Steinernema nematodes, which has emerged as a model system of symbiosis 4. Steinernema nematodes are entomopathogenic, using their bacterial symbiont to kill insects 5. For transmission between insect hosts, the bacteria colonize the intestine of the nematode's infective juvenile stage 6-8. Recently, several other nematode species have been shown to utilize bacteria to kill insects 9-13, and investigations have begun examining the interactions between the nematodes and bacteria in these systems 9.
We describe a method for visualization of a bacterial symbiont within or on a nematode host, taking advantage of the optical transparency of nematodes when viewed by microscopy. The bacteria are engineered to express a fluorescent protein, allowing their visualization by fluorescence microscopy. Many plasmids are available that carry genes encoding proteins that fluoresce at different wavelengths (i.e. green or red), and conjugation of plasmids from a donor Escherichia coli strain into a recipient bacterial symbiont is successful for a broad range of bacteria. The methods described were developed to investigate the association between Steinernema carpocapsae and Xenorhabdus nematophila 14. Similar methods have been used to investigate other nematode-bacterium associations 9,15-18and the approach therefore is generally applicable.
The method allows characterization of bacterial presence and localization within nematodes at different stages of development, providing insights into the nature of the association and the process of colonization 14,16,19. Microscopic analysis reveals both colonization frequency within a population and localization of bacteria to host tissues 14,16,19-21. This is an advantage over other methods of monitoring bacteria within nematode populations, such as sonication 22or grinding 23, which can provide average levels of colonization, but may not, for example, discriminate populations with a high frequency of low symbiont loads from populations with a low frequency of high symbiont loads. Discriminating the frequency and load of colonizing bacteria can be especially important when screening or characterizing bacterial mutants for colonization phenotypes 21,24. Indeed, fluorescence microscopy has been used in high throughput screening of bacterial mutants for defects in colonization 17,18, and is less laborious than other methods, including sonication 22,25-27and individual nematode dissection 28,29.

To study the mutualism between Xenorhabdus bacteria and Steinernema nematodes, methods were developed to monitor bacterial presence and location within nematodes. The experimental approach, which can be applied to other systems, entails engineering bacteria to express the green fluorescent protein and visualizing, using fluorescence microscopy bacteria within the transparent nematode.

 
Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on ...

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

Rapid Analysis and Exploration of Fluorescence Microscopy Images

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized image analysis tools are available, most traditional image-analysis-based workflows have steep learning curves (for fine tuning of analysis parameters) and result in long turnaround times between imaging and analysis. In particular, cell segmentation, the process of identifying individual cells in an image, is a major bottleneck in this regard.
Here we present an alternate, cell-segmentation-free workflow based on PhenoRipper, an open-source software platform designed for the rapid analysis and exploration of microscopy images. The pipeline presented here is optimized for immunofluorescence microscopy images of cell cultures and requires minimal user intervention. Within half an hour, PhenoRipper can analyze data from a typical 96-well experiment and generate image profiles. Users can then visually explore their data, perform quality control on their experiment, ensure response to perturbations and check reproducibility of replicates. This facilitates a rapid feedback cycle between analysis and experiment, which is crucial during assay optimization. This protocol is useful not just as a first pass analysis for quality control, but also may be used as an end-to-end solution, especially for screening. The workflow described here scales to large data sets such as those generated by high-throughput screens, and has been shown to group experimental conditions by phenotype accurately over a wide range of biological systems. The PhenoBrowser interface provides an intuitive framework to explore the phenotypic space and relate image properties to biological annotations. Taken together, the protocol described here will lower the barriers to adopting quantitative analysis of image based screens.

Here we describe a workflow for rapidly analyzing and exploring collections of fluorescence microscopy images using PhenoRipper, a recently developed image-analysis platform.

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized ...

Analysis of Autophagy in Penicillium chrysogenum by Using Starvation Pads in Combination With Fluorescence Microscopy

The study of cellular quality control systems has emerged as a highly dynamic and relevant field of contemporary research. It has become clear that cells possess several lines of defense against damage to biologically relevant molecules like nucleic acids, lipids and proteins. In addition to organelle dynamics (fusion/fission/motility/inheritance) and tightly controlled protease activity, the degradation of surplus, damaged or compromised organelles by autophagy (cellular &#8216;self-eating&#8217;) has received much attention from the scientific community. The regulation of autophagy is quite complex and depends on genetic and environmental factors, many of which have so far not been elucidated. Here a novel method is presented that allows the convenient study of autophagy in the filamentous fungus Penicillium chrysogenum. It is based on growth of the fungus on so-called &#8216;starvation pads&#8217; for stimulation of autophagy in a reproducible manner. Samples are directly assayed by microscopy and evaluated for autophagy induction / progress. The protocol presented here is not limited for use with P. chrysogenum and can be easily adapted for use in other filamentous fungi.

Analysis of Autophagy in Penicillium chrysogenum by Using Starvation Pads in Combination With Fluorescence Microscopy

A convenient and powerful method for studying autophagy in Penicillium chrysogenum by using starvation pads (mixtures of agarose and tap water in a microscope slide containing a central cavity) is presented here.

The study of cellular quality control systems has emerged as a highly dynamic and relevant field of contemporary research. It has become clear that cells ...