Name: simultaneous multicolor imaging of biological structures with fluorescence photoactivation localization microscopy
Uploaded: 2013-12-09T19:00:00
Description: Watch this Scientific Journal Video about Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy at JoVE.com

The authors would like to thank Philip Andresen, Matthew Parent and Sean Carter for computer programming, technical assistance, and useful conversations and Pat Byard for administrative assistance. This work was funded by NIH Career Award K25-AI65459, NIH R15 GM094713, NSF MRI CHE-0722759, Maine Technology Institute MTAF 1106 and 2061, and the Maine Economic Improvement Fund.

Please note: A diagrammatic representation of optical components referenced in this protocol can be found in Figure 1.
1. Cell Sample Preparation
<ol>
	<li>Plate cells at an optimized density (for NIH-3T3 cells, this is roughly 2-5 x 104&#160;cells/cm2) in wells of an 8-well chamber. Cells should be plated in complete media appropriate to the cell type, although media should be made without antibiotics and without phenol red, which contributes to backgr...

<ol>
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Methods. 3, 793-795 (2006).</li><li>Betzig, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+intracellular+fluorescent+proteins+at+nanometer+resolution.">Imaging intracellular fluorescent proteins at nanometer resolution.</a> Sci. 313, 1642-1645 (2006).</li><li>Gould, T. J., Verkhusha, V. V., Hess, S. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+biological+structures+with+fluorescence+photoactivation+localization+microscopy.">Imaging biological structures with fluorescence photoactivation localization microscopy.</a> Nat. Protoc. 4, 291-308 (2009).</li><li>Gould, T. 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U.S.A. 104, 17370-17375 (2007).</li><li>Manley, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-density+mapping+of+single-molecule+trajectories+with+photoactivated+localization+microscopy.">High-density mapping of single-molecule trajectories with photoactivated localization microscopy.</a> Nat. Methods. 5, 155-157 (2008).</li><li>Shroff, H., Galbraith, C. G., Galbraith, J. A., Betzig, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live-cell+photoactivated+localization+microscopy+of+nanoscale+adhesion+dynamics.">Live-cell photoactivated localization microscopy of nanoscale adhesion dynamics.</a> Nat. 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Localization-based super-resolution imaging provides many powerful capabilities for biological imaging. The route from individual optical components placed on the table to a functional super-resolution microscope capable of simultaneously imaging multiple fluorescent species in a biological sample presents a number of challenges. Some aspects of the alignment are more critical than others; we endeavor below to provide guidance to prospective users dealing with the most difficult aspects of the route.
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While cellular structures exist on a wide range of spatial scales, fluorescence imaging of cellular organization on length scales smaller than ~250 nm is restricted in conventional microscopy due to the physical constraint of the diffraction limit. This limit was overcome with the advent of fluorescence photoactivation localization microscopy (FPALM1) and similar techniques2,3,&#160;which can localize large numbers of individual molecules with precision of ~10 nm, to generate images with resolution of a few tens of nanometers. FPALM is based on using optical control to activate and inactivate subsets of molecules (for a full description of FPALM, and instructions on how to implement this imaging system, see Gould et al.4). This technique allows for the spatial distributions of whole populations of single molecules to be mapped, thereby elucidating biological structures across length scales spanning from tens of nanometers to tens of microns. Localization-based super-resolution microscopy (hereto referred to as localization microscopy) has now been adapted to address a range of biological questions, with technological developments permitting, for example, the imaging of individual molecular orientations with polarization FPALM, or P-FPALM5,&#160;the fluorescence imaging of single molecules in three dimensions with Biplane FPALM6&#160;or other techniques7-9,&#160;and the super-resolution fluorescence imaging of single molecules in living cells10-12.&#160;Localization microscopy has also been applied to the imaging of multiple species in fixed cells13-16.&#160;Recently, three protein species have been simultaneously imaged with FPALM in both fixed and living cells17.&#160;Localization microscopy can image samples labeled in a variety of ways: examples include proteins expressed with PAFP or PSFP fusion tags, antibodies or molecules labeled with caged organic dyes, or conventional organic dyes. While the use of conventional fluorescent dyes allows for the labeling of proteins in the absence of a fusion-protein tag, the conditions generally required for the use of noncaged organic dyes in super-resolution imaging require samples to be immersed in reducing buffers2.&#160;Additionally, the intracellular delivery of antibody-dye conjugates typically requires cells to be fixed and their membranes permeabilized, or requires that living cells are made permeable through electroporation or some other means. The requirements for reducing buffer conditions and membrane permeabilization limit the suitability of organic dyes for live cell imaging, although recent developments have allowed for effective use of HaloTags and FPALM to image membrane structures18.
FPALM was the first localization microscopy technique to be applied to live cells10.&#160;In live cells, in addition to providing a time dependent spatial map of the locations of labeled molecules, FPALM can track single molecules over multiple frames, and molecular trajectories determined over timescales of milliseconds19.&#160;Thus, FPALM provides access to fairly short timescales and nanoscale resolution.
Multicolor FPALM can be used for a variety of different probes, including photoactivatable proteins and organic caged or noncaged dyes. We here provide detail on the protocol and setup for the simultaneous imaging of two fluorescent protein species, Dendra2 and PAmCherry. We report the outcomes of imaging PAmCherry conjugated to beta actin (PAmCherry-actin) and Dendra2 conjugated to influenza hemagglutinin (Dendra2-HA) in NIH-3T3 fibroblasts. Components described in the setup can be interchanged for other hardware more suited to the imaging of other probes. Where this is the case, we have tried to be explicit in the text.
Multicolor FPALM is ideal for reporting the spatial distributions of multiple protein species in living or fixed cells. This technique is especially suited to investigating spatial and/or dynamic relationships on nanometer length spatial scales, although images will report localization on a range of length scales, from tens of nanometers up to tens of microns. One major advantage of multicolor FPALM is that the setup is relatively inexpensive to construct, and very flexible for use with various probe combinations. The process of construction and calibration of the system from components also provides considerable understanding of factors which can compromise the quality and interpretability of the data, and so the research outcome. We here detail the methods for the optical setup, sample preparation, and data acquisition of multiple protein species, with PSFP and PAFP fusion constructs, using FPALM. While this protocol describes the analysis of fixed cells, these procedures are readily applicable to the imaging of living cells.
The optical setup here described is ideal for the simultaneous imaging of the PSFP Dendra2 and the PAFP PAmCherry. Many other probes may be used for multicolor imaging; however, the precise components required may vary, depending on the excitation and emission spectra of the chosen probes. Choices of dichroic mirrors, filters, and laser wavelengths should be made based on these considerations.

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		<col />
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	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:195px;">LabTek II chambers</td>
			<td style="width:101px;">Nunc</td>
			<td style="width:137px;"> </td>
			<td style="width:183px;"> </td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:195px;">Fluorescent beads</td>
			<td style="width:101px;">Invitrogen</td>
			<td style="width:137px;">F-8801</td>
			<td style="width:183px;">Beads for calibration</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:195px;">Tetraspeck beads</td>
			<td style="width:101px;">Invitrogen</td>
			<td style="width:137px;">T-7279</td>
			<td style="width:183px;">Four color beads for calibration</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:195px;">Objective immersion oil</td>
			<td style="width:101px;">Zeiss</td>
			<td style="width:137px;">518F</td>
			<td style="width:183px;">Immersion oil for high NA objective (dependent on choice of objective)</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:195px;">HPLC water</td>
			<td style="width:101px;">Fisher Scientific</td>
			<td style="width:137px;">W5-4</td>
			<td style="width:183px;"> </td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:195px;">Media</td>
			<td style="width:101px;">ATCC</td>
			<td style="width:137px;">30-2003</td>
			<td style="width:183px;">Or Cellgro 10-090</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:195px;">Antibiotics</td>
			<td style="width:101px;">GIBCO</td>
			<td style="width:137px;">15070-063</td>
			<td style="width:183px;"> </td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:195px;">serum</td>
			<td style="width:101px;">Thermo Scientific</td>
			<td style="width:137px;">SH30087.03</td>
			<td style="width:183px;"> </td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:195px;">Lipofectamine</td>
			<td style="width:101px;">Invitrogen</td>
			<td align="right" style="width:137px;">52887</td>
			<td style="width:183px;"> </td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:195px;">Optimem I</td>
			<td style="width:101px;">GIBCO</td>
			<td style="width:137px;">11058-021</td>
			<td style="width:183px;"> </td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:195px;">Trypsin</td>
			<td style="width:101px;">MPBiomedicals</td>
			<td align="right" style="width:137px;">1689149</td>
			<td style="width:183px;"> </td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:195px;">paraformaldehyde</td>
			<td style="width:101px;">Fisher Scientific</td>
			<td style="width:137px;">AA433689M</td>
			<td style="width:183px;">CAUTION: Toxic</td>
		</tr>
	</tbody>
</table>

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.

Influenza hemagglutinin (HA) forms clusters on the order of tens of nanometers to micrometers, and these clusters variably colocalize with actin (Figure 5). These spatial distributions corroborate coarser scale imaging of these two proteins28, and the dependence of the HA spatial distributions on actin19.&#160;Multicolor FPALM images can be further used to describe the density, area and perimeter of these clusters, and the degree of colocalization between the two species at both the...

Watch this Scientific Journal Video about Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy at JoVE.com

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

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 ...

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Biology

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 ...

Differential Imaging of Biological Structures with Doubly-resonant Coherent Anti-stokes Raman Scattering (CARS)

Coherent Raman imaging techniques have seen a dramatic increase in activity over the past decade due to their promise to enable label-free optical imaging with high molecular specificity 1. The sensitivity of these techniques, however, is many orders of magnitude weaker than fluorescence, requiring milli-molar molecular concentrations 1,2. Here, we describe a technique that can enable the detection of weak or low concentrations of Raman-active molecules by amplifying their signal with that obtained from strong or abundant Raman scatterers. The interaction of short pulsed lasers in a biological sample generates a variety of coherent Raman scattering signals, each of which carry unique chemical information about the sample. Typically, only one of these signals, e.g. Coherent Anti-stokes Raman scattering (CARS), is used to generate an image while the others are discarded. However, when these other signals, including 3-color CARS and four-wave mixing (FWM), are collected and compared to the CARS signal, otherwise difficult to detect information can be extracted 3. For example, doubly-resonant CARS (DR-CARS) is the result of the constructive interference between two resonant signals 4. We demonstrate how tuning of the three lasers required to produce DR-CARS signals to the 2845 cm-1 CH stretch vibration in lipids and the 2120 cm-1 CD stretching vibration of a deuterated molecule (e.g. deuterated sugars, fatty acids, etc.) can be utilized to probe both Raman resonances simultaneously. Under these conditions, in addition to CARS signals from each resonance, a combined DR-CARS signal probing both is also generated. We demonstrate how detecting the difference between the DR-CARS signal and the amplifying signal from an abundant molecule's vibration can be used to enhance the sensitivity for the weaker signal. We further demonstrate that this approach even extends to applications where both signals are generated from different molecules, such that e.g. using the strong Raman signal of a solvent can enhance the weak Raman signal of a dilute solute.

Differential Imaging of Biological Structures with Doubly-resonant Coherent Anti-stokes Raman Scattering CARS

A combination of three single wavelength short-pulsed lasers is used to generate coherent anti-Stokes Raman scattering (CARS) and doubly-resonant CARS (DR-CARS). The difference between these signals provides enhanced sensitivity for otherwise difficult to detect coherent Raman signals, enabling imaging of weak Raman scatterers.

Coherent Raman imaging techniques have seen a dramatic increase in activity over the past decade due to their promise to enable label-free optical imaging ...

Nano-fEM: Protein Localization Using Photo-activated Localization Microscopy and Electron Microscopy

Mapping the distribution of proteins is essential for understanding the function of proteins in a cell. Fluorescence microscopy is extensively used for protein localization, but subcellular context is often absent in fluorescence images. Immuno-electron microscopy, on the other hand, can localize proteins, but the technique is limited by a lack of compatible antibodies, poor preservation of morphology and because most antigens are not exposed to the specimen surface. Correlative approaches can acquire the fluorescence image from a whole cell first, either from immuno-fluorescence or genetically tagged proteins. The sample is then fixed and embedded for electron microscopy, and the images are correlated 1-3. However, the low-resolution fluorescence image and the lack of fiducial markers preclude the precise localization of proteins. 
Alternatively, fluorescence imaging can be done after preserving the specimen in plastic. In this approach, the block is sectioned, and fluorescence images and electron micrographs of the same section are correlated 4-7. However, the diffraction limit of light in the correlated image obscures the locations of individual molecules, and the fluorescence often extends beyond the boundary of the cell. 
Nano-resolution fluorescence electron microscopy (nano-fEM) is designed to localize proteins at nano-scale by imaging the same sections using photo-activated localization microscopy (PALM) and electron microscopy. PALM overcomes the diffraction limit by imaging individual fluorescent proteins and subsequently mapping the centroid of each fluorescent spot 8-10. 
We outline the nano-fEM technique in five steps. First, the sample is fixed and embedded using conditions that preserve the fluorescence of tagged proteins. Second, the resin blocks are sectioned into ultrathin segments (70-80 nm) that are mounted on a cover glass. Third, fluorescence is imaged in these sections using the Zeiss PALM microscope. Fourth, electron dense structures are imaged in these same sections using a scanning electron microscope. Fifth, the fluorescence and electron micrographs are aligned using gold particles as fiducial markers. In summary, the subcellular localization of fluorescently tagged proteins can be determined at nanometer resolution in approximately one week.

We describe a method to localize fluorescently tagged proteins in electron micrographs. Fluorescence is first localized using photo-activated localization microscopy on ultrathin sections. These images are then aligned to electron micrographs of the same section.

Mapping the distribution of proteins is essential for understanding the function of proteins in a cell. Fluorescence microscopy is extensively used for ...

Analyzing and Building Nucleic Acid Structures with 3DNA

The 3DNA software package is a popular and versatile bioinformatics tool with capabilities to analyze, construct, and visualize three-dimensional nucleic acid structures. This article presents detailed protocols for a subset of new and popular features available in 3DNA, applicable to both individual structures and ensembles of related structures. Protocol 1 lists the set of instructions needed to download and install the software. This is followed, in Protocol 2, by the analysis of a nucleic acid structure, including the assignment of base pairs and the determination of rigid-body parameters that describe the structure and, in Protocol 3, by a description of the reconstruction of an atomic model of a structure from its rigid-body parameters. The most recent version of 3DNA, version 2.1, has new features for the analysis and manipulation of ensembles of structures, such as those deduced from nuclear magnetic resonance (NMR) measurements and molecular dynamic (MD) simulations; these features are presented in Protocols 4 and 5. In addition to the 3DNA stand-alone software package, the w3DNA web server, located at <a href="http://w3dna.rutgers.edu" target="_blank">http://w3dna.rutgers.edu</a>, provides a user-friendly interface to selected features of the software. Protocol 6 demonstrates a novel feature of the site for building models of long DNA molecules decorated with bound proteins at user-specified locations.

The 3DNA software package is a popular and versatile bioinformatics tool with capabilities to analyze, construct, and visualize three-dimensional nucleic acid structures. This article presents detailed protocols for a subset of new and popular features available in 3DNA, applicable to both individual structures and ensembles of related structures.

The 3DNA software package is a popular and versatile bioinformatics tool with capabilities to analyze, construct, and visualize three-dimensional nucleic ...

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...

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 ...

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.

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 ...

Quantitative Localization of a Golgi Protein by Imaging Its Center of Fluorescence Mass

The Golgi complex consists of serially stacked membrane cisternae which can be further categorized into sub-Golgi regions, including the cis-Golgi, medial-Golgi, trans-Golgi and trans-Golgi network. Cellular functions of the Golgi are determined by the characteristic distribution of its resident proteins. The spatial resolution of conventional light microscopy is too low to resolve sub-Golgi structure or cisternae. Thus, the immuno-gold electron microscopy is a method of choice to localize a protein at the sub-Golgi level. However, the technique and instrument are beyond the capability of most cell biology labs. We describe here our recently developed super-resolution method called Golgi protein localization by imaging centers of mass (GLIM) to systematically and quantitatively localize a Golgi protein. GLIM is based on standard fluorescence labeling protocols and conventional wide-field or confocal microscopes. It involves the calibration of chromatic-shift aberration of the microscopic system, the image acquisition and the post-acquisition analysis. The sub-Golgi localization of a test protein is quantitatively expressed as the localization quotient. There are four main advantages of GLIM; it is rapid, based on conventional methods and tools, the localization result is quantitative, and it affords ~ 30 nm practical resolution along the Golgi axis. Here we describe the detailed protocol of GLIM to localize a test Golgi protein.

The precise localization of Golgi residents is essential for understanding the cellular functions of the Golgi. However, conventional optical microscopy is unable to resolve the sub-Golgi structure. Here we describe the protocol for a conventional microscopy based super-resolution method to quantitatively determine the sub-Golgi localization of a protein.

The Golgi complex consists of serially stacked membrane cisternae which can be further categorized into sub-Golgi regions, including the cis-Golgi, ...