Name: live-cell fluorescence microscopy to investigate subcellular protein localization and cell morphology changes in bacteria
Uploaded: 2019-11-23T14:00:00.000+00:00
Duration: 5 min 57 s
Description: Watch this Scientific Journal Video about Live-Cell Fluorescence Microscopy to Investigate Subcellular Protein Localization and Cell Morphology Changes in Bacteria at JoVE.com

We thank our lab members for their comments on this article. This work was funded by a start-up grant from the University of South Florida (PE).

1. General growth conditions
<ol>
	<li>Inoculate 2 mL of appropriate growth medium supplemented with antibiotics (where required) with a single colony of the strain(s) to be imaged. Incubate these seed cultures overnight at 22 &#176;C in a shaking incubator. 
	NOTE: The specific bacterial growth conditions used in this article are provided under the representative results section.</li>
	<li>Dilute overnight cultures 1:20 in fresh media in a 125 mL flask, supplement with antibiotics (where required).</li>
	<li>Grow culture(s) at 37 &#176;C in a shaking incubator until mid-logarithmic phase (OD600 = 0.5). 
	NOTE: Inducer or inhibitor could be added directly to growing culture(s) at the appropriate growth phase.</li>
	<li>Harvest cells at desired culture conditions for microscopy sample preparation as described in the following section.</li>
</ol>
2. Sample preparation
<ol>
	<li>Prepare 1% agarose by combining 0.25 g of molecular biology grade, low EEO, agarose with either 25 mL of growth medium supplemented with appropriate antibiotics (where required) for timelapse microscopy or sterile water. Heat the mixture with the help of a microwave for approximately 30 s and pour it into a sterile 100 mm x 15 mm Petri dish and let it solidify.</li>
	<li>Take a 5&#8211;50 &#181;L aliquot of the cell culture depending on the need and stain the cells. Stain cells with appropriate dyes at this stage (e.g., add fluorescent dye FM4-64 (membrane stain) for an experiment (Figure 1)).
	<ol>
		<li>Pipette a 5 &#181;L aliquot of the culture sample or stained sample to be imaged onto the bottom of a 35 mm glass bottom culture dish (with 14 mm microwell diameter and uncoated No. 1.5 coverslip; see the Table of Materials). 
		CAUTION: Do not use poly-L-lysine coated coverslips especially for timelapse microscopy as they could induce different growth pattern (we have observed this with poly-D-lysine as well; data not shown) and/or affect protein dynamics6.</li>
		<li>To minimize the use of dyes, stain 3&#8211;4 &#181;L cell suspension (harvested at OD600 = 0.5; approximately 2.7 x 107 and 3.4 x 107 for B. subtilis and S. aureus respectively per 1 mL culture) directly on the glass bottom dish.</li>
		<li>Place a pre-cut agarose slab (11 mm in diameter or of any desired size to overlap the area of the coverslip; cut using the open end of a sterile tube or a razor blade) on top of the sample and gently tap to make sure the agarose slab is lying flat against the coverslip.</li>
	</ol>
	</li>
	<li>Subsequent to imaging (see following section), if the number of cells in the field of view are not desirable, adjust the cell density of the sample via dilution or concentration by centrifugation and alter the ratio of cells in the sample using growth medium/buffer as necessary. 
	NOTE: To minimize autofluorescence emanating from the culture medium, cells can be washed in buffer (for example in standard 1x phosphate buffer saline) and resuspended in appropriate amount of buffer prior to imaging. It is possible that in this step smaller cells or vesicles may not be retained after centrifugation and will hence be lost.</li>
	<li>Add water using a pipette (~5 &#181;L drops) to the space inside the culture dish (around the coverslip) to prevent the agarose pad from drying, and to help maintain humidity during the course of image acquisition, especially for timelapse experiments.</li>
	<li>Allow culture dishes to equilibrate to the temperature inside the incubation chamber, a built-in opaque compartment provided by the manufacturer, of the microscope for 15&#8211;20 min. 
	NOTE: Turn on the heating element and set the incubation chamber to a desired temperature several hours prior to imaging. This will ensure that the hardware stabilizes to the new temperature. For prolonged timelapse imaging, place a beaker or flask with water inside the microscope chamber, away from the working area, to maintain humidity.</li>
</ol>
3. Imaging
<ol>
	<li>On the day of the experiment, turn on the microscope system. Start the imaging software (see Table of Materials) by clicking the appropriate icon on the desktop. Initialize the microscope by clicking the Initialize Microscope option (button depicted with a microscope on it) on the software&#8217;s start-up dialog box. Ensure that the objective is fully lowered using the microscope coarse adjustment prior to initialization. 
	NOTE: Following initialization three additional dialog boxes should appear in addition to the start menu: resolve3D, data collection, and filter monitor.</li>
	<li>Place a drop of 1.517 (refractive index) oil on the 100x oil immersion objective supplied by the manufacturer (Numerical Aperture = 1.4, Working Distance = 0.12 mm; see Table of Materials). 
	NOTE: It is important to choose the appropriate immersion oil for the temperature at which imaging is conducted.</li>
	<li>Load the glass bottom dish containing sample into the metal housing (coffin) and gently slide into the stage clamp.</li>
	<li>Use the coarse adjustment knob to raise the objective until the oil makes contact with the glass bottom of the dish. Use the eye piece and fine adjustment knob to bring the sample into focus. Once the cells are in focus turn the knob from the eye piece to camera mode by moving the selector switch located on the front of the microscope body to the left.</li>
	<li>Begin experiment using the imaging software. On the resolve3D window, select the Design/run experiment icon depicted by a flask. A new dialog box should appear entitled design/run experiment.
	<ol>
		<li>Set the number of Z-stacks and sample thickness using the design and then sectioning tab on the design/run experiment dialog box. 
		NOTE: For the experiments in representative results section, 17 Z-stacks at 200 nm interval for still images and four Z-stacks at 200 nm interval for timelapse microscopy were used.</li>
		<li>To measure the thickness of the cells in the sample, manually adjust the Z-plane incrementally by using the up and down arrows on the resolve3D dialog box. Mark where the cells go out-of-focus as the upper and lower limit for image acquisition. Import this information prior to running the experiment.</li>
		<li>To help minimize phototoxicity and photobleaching during timelapse imaging, reduce the number of Z-stacks and choose the mid-plane of the cells for image acquisition.</li>
	</ol>
	</li>
	<li>Select the appropriate filter set for the experiment using the design and then channels tab on the design/run experiment dialog box (TRITC: EX 542/27; EM 597/45; FITC/GFP: EX 475/28; EM 525/48; mCherry: EX 575/25; EM 632/60; Cy5: EX 632/22; EM 676/34).
	<ol>
		<li>Also, select a reference for the collection of POL/DIC information. Adjust percentage transmission (light intensity) and duration of exposure for individual channels selected prior to imaging by selecting the appropriate options on the resolve3D dialog box. 
		NOTE: In a test field of view that is not considered for the experiment test these settings to identify if the selected parameters obtain meaningful fluorescence data without missing weak signal or oversaturating it. Then import these parameters for the experimental set up.</li>
	</ol>
	</li>
	<li>Open the points list by selecting the Points list button on the resolve3D dialog box. A new dialog box should appear entitled points list. Mark several fields of view to be used in the experiment by finding appropriate fields of view using the microscope stage controls and selecting the mark point option on the points list dialog box. 
	NOTE: A replace point will have to be selected each time the microscope is refocused on a point in the points list. This can be done by focusing the microscope on the appropriate point in the list and then selecting the replace point button. It is important to not touch the analog coarse/fine adjustment knob when refocusing; use only the software to adjust the focus.</li>
	<li>Set timelapse parameters by first selecting the design and then timelapse tab on the design/run experiment dialog box. Select the timelapse check box. Enter the appropriate timelapse parameters in terms of timelapse images/total time.</li>
	<li>Set points to be imaged from the points list by first selecting the design and then points tab on the design/run experiment dialog box. Select the visit points list option and enter points to be imaged in the text box separated by commas or hyphens if it is a complete sequence.</li>
	<li>Prior to beginning an experiment, edit file names and file locations using the run tab on the design/run dialog box. File location can be changed by selecting the settings button, selecting the data folder, and by then selecting the appropriate folder or creating a new one. Change file names by entering in the new file name in the image file name text box.</li>
	<li>Begin the experiment by selecting the play button on the begin experiment dialog box. 
	NOTE: For timelapse, continually check focus at each field of view throughout the experiment using DIC setting (to avoid unnecessary photobleaching) and refocus and update the information in points list as cells tend to go out of focus over time.</li>
</ol>
4. Image processing
<ol>
	<li>Open the desired raw (R3D) image files for generation of figures. 
	NOTE: Recently saved files can be located using the data folder button on the imaging software&#8217;s start-up dialog box.</li>
	<li>Run the deconvolution program, to remove out-of-focus fluorescence light7,8, in order to produce a deconvolved D3D image file. Select the process tab on the start-up dialog box, and then select deconvolve. A new dialog box will appear entitled deconvolve.
	<ol>
		<li>Set the deconvolution parameters by dragging the appropriate image number (located in the upper left-hand corner of each image file) to the input on the deconvolution dialog box. On the deconvolution dialog box, select the more options button, and deselect the crop border rolloff after processing check box (otherwise the images cannot be superimposed over the corresponding DIC file).</li>
		<li>Click the do it button on the deconvolution dialog box to deconvolve raw image file. 
		NOTE: The deconvolution parameters could be adjusted as indicated by the software manufacturer&#8217;s guidance manual for desired results.</li>
	</ol>
	</li>
	<li>If necessary, perform manual background noise subtraction and brightness/contrast adjustment in any or all wavelength (color) channels. Do this by selecting the contrast adjustment button on the deconvolved image file.</li>
	<li>Save the image as a TIFF file by first selecting the file option at the top of the D3D dialog box. Then select save as TIFF, identify and select appropriate Z-stacks (that are within focus), select or unselect desired filter sets.</li>
	<li>Perform data quantification using any manufacturer-supplied software or freely available programs such as ImageJ5.
	<ol>
		<li>For cell size quantification click on the tool tab on the appropriate D3D image file, and then select the measure distances option. Measure distances by selecting start and end points on the desired image file using the mouse through left clicks. 
		NOTE: It is best to zoom in on the image during cell size quantification.</li>
		<li>For fluorescence signal quantification use the data inspector option under the tool tab on the appropriate D3D image file. 
		NOTE: It is best to zoom in on the image and to have only relevant filters selected when completing fluorescence signal quantification.</li>
		<li>To quantify the fluorescence signal, draw a box with column/row option set to specific dimension. Select an area with signal to be quantified and record the value. Measure the background signal by using the same size box to select an area immediately outside of the cells. Subtract the background value from the value recorded from the fluorescence signal obtained within the cell.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Microscopy has remained a mainstay in studies pertaining to microbial organisms. Given their micron-scale cell size, single-cell level studies have traditionally relied on electron microscopy (EM). Although EM has become quite a powerful technique in recent years, it has its own intrinsic limitations in addition to limited user access16. Improvements in fluorescence microscopy techniques and development of different fluorescent probes, such as FDAA3, have provided microbial...

The field of bacterial cell biology has been significantly enhanced by recent advancements in microscopy techniques1,2. Among other instruments, microscopes that are capable of conducting timelapse fluorescence microscopy experiments remain a valuable tool. Investigators can monitor various physiological events in real-time using fluorescent proteins such as, green fluorescent protein (GFP)-based transcriptional and translational reporter fusions, fluorescent D-amino acids (FDAA)3, or use other stains for labeling the cell wall, membrane and DNA. It is therefore of no surprise that fluorescence microscopy remains popular among microbial cell biologists. In addition to simply showing the end phenotypes, providing information as to how the observed phenotypes arise using timelapse microscopy could add significant value to the findings and potentially offer clues as to what cellular processes are being targeted by potential drug candidates4.
The protocols to conduct high-resolution imaging using a fully motorized, inverted, wide-field fluorescence microscope (see the Table of Materials) are provided in this article. These protocols could be adapted to suit the needs of other fluorescence microscopes that are capable of conducting timelapse microscopy. Although the software discussed here corresponds to the specific manufacturer-supplied software as indicated in the Table of Materials, software commonly supplied by other microscope manufacturers or the freely available ImageJ5, have equivalent tools for analyzing microscopy data. For conditions where timelapse is not conducive, time-course experiments could be conducted as described in this article. The protocols described here provide a detailed guide to study the phenotypic changes in two different bacterial species: B. subtilis and S. aureus. See Table 1 for strains used.

<table><tbody><tr><td>Agarose</td><td>Fisher BioReagents</td><td>BP160-100</td><td>Molecular Biology Grade - Low EEO</td></tr><tr><td>DAPI</td><td>Invitrogen</td><td>D3571</td><td>Microscopy</td></tr><tr><td>FM4-64</td><td>Invitrogen</td><td>T3166</td><td>Microscopy</td></tr><tr><td>Glass bottom dish</td><td>MatTek</td><td>P35G-1.5-14-C</td><td>Microscopy</td></tr><tr><td>IPTG</td><td>Fisher BioReagents</td><td>BP1755-10</td><td>Dioxane-free</td></tr><tr><td>Microscope</td><td>GE</td><td>DeltaVision Elite</td><td>Customized Olympus IX-71 Inverted Microscope Stand; Custom Illumination Tower and Transmitted Light Illuminator Module. Objectives: PLAPON 60X (N.A. 1.42, WD 0.15 mm); OLY 100X OIL (N.A. 1.4, WD 0.12 mm); DIC Prism Nomarski for 100X Objective; CoolSnap HQ2 camera; SSI Assembly 7-color; Environmental control chamber - opaque.</td></tr><tr><td>PC190723</td><td>MilliporeSigma</td><td>3445805MG</td><td>FtsZ inhibitor</td></tr><tr><td>SoftWorx</td><td>GE</td><td></td><td>Manufacturer-supplied imaging software</td></tr></tbody></table>

live-cell fluorescence microscopy to investigate subcellular protein localization and cell morphology changes in bacteria

Investigations of factors influencing cell division and cell shape in bacteria are commonly performed in conjunction with high-resolution fluorescence microscopy as observations made at a population level may not truly reflect what occurs at a single cell level. Live-cell timelapse microscopy allows investigators to monitor the changes in cell division or cell morphology which provide valuable insights regarding subcellular localization of proteins and timing of gene expression, as it happens, to potentially aid in answering important biological questions. Here, we describe our protocol to monitor phenotypic changes in Bacillus subtilis and Staphylococcus aureus using a high-resolution deconvolution microscope. The objective of this report is to provide a simple and clear protocol that can be adopted by other investigators interested in conducting fluorescence microscopy experiments to study different biological processes in bacteria as well as other organisms.

GpsB phenotypes 
Previously we have shown that Sa-GpsB is an essential protein as depletion of GpsB using an antisense RNA results in cell lysis9. Here we describe how the emergence of various cell division phenotypes and changes in protein localization could be captured using the timelapse microscopy protocol described in this article. For this purpose, S. aureus strains RB143 [SH1000 harboring pEPSA5 (empty vector)] and GGS8 [SH1000 harboring pGG59 (P...

Watch this Scientific Journal Video about Live-Cell Fluorescence Microscopy to Investigate Subcellular Protein Localization and Cell Morphology Changes in Bacteria at JoVE.com

Live-Cell Fluorescence Microscopy to Investigate Subcellular Protein Localization and Cell Morphology Changes in Bacteria

This article provides a step-by-step guide to investigate protein subcellular localization dynamics and to monitor morphological changes using high-resolution fluorescence microscopy in Bacillus subtilis and Staphylococcus aureus.

Investigations of factors influencing cell division and cell shape in bacteria are commonly performed in conjunction with high-resolution fluorescence ...

live-cell-fluorescence-microscopy-to-investigate-subcellular-protein

Research

JoVE Journal

Immunology and Infection

6.9K Views.  University of South Florida. This article provides a step-by-step guide to investigate protein subcellular localization dynamics and to monitor morphological changes using high-resolution fluorescence microscopy in Bacillus subtilis and Staphylococcus aureus.

Video: Live-Cell Fluorescence Microscopy to Investigate Subcellular Protein Localization and Cell Morphology Changes in Bacteria

Monitoring Spatial Segregation in Surface Colonizing Microbial Populations

Microbes provide an intriguing system to study social interaction among individuals within a population. The short generation times and relatively simple genetic modification procedures of microbes facilitate the development of the sociomicrobiology field. To assess the fitness of certain microbial species, selected strains or their genetically modified derivatives within one population, can be fluorescently labelled and tracked using microscopy adapted with appropriate fluorescence filters. Expanding colonies of diverse microbial species on agar media can be used to monitor the spatial distribution of cells producing distinctive fluorescent proteins.
Here, we present a detailed protocol for the use of green- and red-fluorescent protein producing bacterial strains to follow spatial arrangement during surface colonization, including flagellum-driven community movement (swarming), exopolysaccharide- and hydrophobin-dependent growth mediated spreading (sliding), and complex colony biofilm formation. Non-domesticated isolates of the Gram-positive bacterium, Bacillus subtilis can be utilized to scrutinize certain surface spreading traits and their effect on two-dimensional distribution on the agar-solidified medium. By altering the number of cells used to initiate colony biofilms, the assortment levels can be varied on a continuous scale. Time-lapse fluorescent microscopy can be used to witness the interaction between different phenotypes and genotypes at a certain assortment level and to determine the relative success of either.

The role of assortment (spatial segregation) in evolutionary scenarios can be examined using simple microbial systems in the laboratory that allow controlled adjustment of spatial distribution. By modifying the founder cell density, various assortment levels can be visualized using fluorescently labelled bacterial strains in the colony biofilms of Bacillus subtilis.

Microbes provide an intriguing system to study social interaction among individuals within a population. The short generation times and relatively simple ...

Genetics

Live Cell Imaging of Bacillus subtilis and Streptococcus pneumoniae using Automated Time-lapse Microscopy

During the last few years scientists became increasingly aware that average data obtained from microbial population based experiments are not representative of the behavior, status or phenotype of single cells. Due to this new insight the number of single cell studies rises continuously (for recent reviews see 1,2,3). However, many of the single cell techniques applied do not allow monitoring the development and behavior of one specific single cell in time (e.g. flow cytometry or standard microscopy).
Here, we provide a detailed description of a microscopy method used in several recent studies 4, 5, 6, 7, which allows following and recording (fluorescence of) individual bacterial cells of Bacillus subtilis and Streptococcus pneumoniae through growth and division for many generations. The resulting movies can be used to construct phylogenetic lineage trees by tracing back the history of a single cell within a population that originated from one common ancestor. This time-lapse fluorescence microscopy method cannot only be used to investigate growth, division and differentiation of individual cells, but also to analyze the effect of cell history and ancestry on specific cellular behavior. Furthermore, time-lapse microscopy is ideally suited to examine gene expression dynamics and protein localization during the bacterial cell cycle. The method explains how to prepare the bacterial cells and construct the microscope slide to enable the outgrowth of single cells into a microcolony. In short, single cells are spotted on a semi-solid surface consisting of growth medium supplemented with agarose on which they grow and divide under a fluorescence microscope within a temperature controlled environmental chamber. Images are captured at specific intervals and are later analyzed using the open source software ImageJ.

Live Cell Imaging of Bacillus subtilis and Streptococcus pneumoniae using Automated Time-lapse Microscopy

This protocol provides a step-by-step procedure to monitor single cell behavior of different bacteria in time using automated fluorescence time-lapse microscopy. Furthermore, we provide guidelines how to analyze the microscopy images.

During the last few years scientists became increasingly aware that average data obtained from microbial population based experiments are not ...

Single-cell Analysis of Bacillus subtilis Biofilms Using Fluorescence Microscopy and Flow Cytometry

Biofilm formation is a general attribute to almost all bacteria 1-6. When bacteria form biofilms, cells are encased in extracellular matrix that is mostly constituted by proteins and exopolysaccharides, among other factors 7-10. The microbial community encased within the biofilm often shows the differentiation of distinct subpopulation of specialized cells 11-17. These subpopulations coexist and often show spatial and temporal organization within the biofilm 18-21.
Biofilm formation in the model organism Bacillus subtilis requires the differentiation of distinct subpopulations of specialized cells. Among them, the subpopulation of matrix producers, responsible to produce and secrete the extracellular matrix of the biofilm is essential for biofilm formation 11,19. Hence, differentiation of matrix producers is a hallmark of biofilm formation in B. subtilis.
We have used fluorescent reporters to visualize and quantify the subpopulation of matrix producers in biofilms of B. subtilis 15,19,22-24. Concretely, we have observed that the subpopulation of matrix producers differentiates in response to the presence of self-produced extracellular signal surfactin 25. Interestingly, surfactin is produced by a subpopulation of specialized cells different from the subpopulation of matrix producers 15.
We have detailed in this report the technical approach necessary to visualize and quantify the subpopulation of matrix producers and surfactin producers within the biofilms of B. subtilis. To do this, fluorescent reporters of genes required for matrix production and surfactin production are inserted into the chromosome of B. subtilis. Reporters are expressed only in a subpopulation of specialized cells. Then, the subpopulations can be monitored using fluorescence microscopy and flow cytometry (See Fig 1).
The fact that different subpopulations of specialized cells coexist within multicellular communities of bacteria gives us a different perspective about the regulation of gene expression in prokaryotes. This protocol addresses this phenomenon experimentally and it can be easily adapted to any other working model, to elucidate the molecular mechanisms underlying phenotypic heterogeneity within a microbial community.

Single-cell Analysis of Bacillus subtilis Biofilms Using Fluorescence Microscopy and Flow Cytometry

Microbial biofilms are generally constituted by distinct subpopulations of specialized cells. Single-cell analysis of these subpopulations requires the use of fluorescent reporters. Here we describe a protocol to visualize and monitor several subpopulationswithin B. subtilis biofilms using fluorescence microscopy and flow cytometry.

Biofilm formation is a general attribute to almost all bacteria 1-6. When bacteria form biofilms, cells are encased in extracellular matrix that is mostly ...

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

The obligate intracellular bacterium Chlamydia elicits a great burden on global public health. C.&#160;trachomatis is the leading bacterial cause of sexually transmitted infection and also the primary cause of preventable blindness in the world. An essential determinant for successful infection of host cells by Chlamydia is the bacterium's ability to manipulate host cell signaling from within a novel, vacuolar compartment called the inclusion. From within the inclusion, Chlamydia acquire nutrients required for their 2-3 day developmental growth, and they additionally secrete a panel of effector proteins onto the cytosolic face of the vacuole membrane and into the host cytosol. Gaps in our understanding of Chlamydia biology, however, present significant challenges for visualizing and analyzing this intracellular compartment. Recently, a reverse-imaging strategy for visualizing the inclusion using GFP expressing host cells was described. This approach rationally exploits the intrinsic impermeability of the inclusion membrane to large molecules such as GFP. In this work, we describe how GFP- or mCherry-expressing host cells are generated for subsequent visualization of chlamydial inclusions. Furthermore, this method is shown to effectively substitute for costly antibody-based enumeration methods, can be used in tandem with other fluorescent labels, such as GFP-expressing Chlamydia, and can be exploited to derive key quantitative data about inclusion membrane growth from a range of Chlamydia species and strains.

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

A live cell fluorescent protein based method for illuminating cellular vacuoles (inclusions) containing Chlamydia is described. This strategy enables rapid, automated determination of Chlamydia infectivity in samples and can be used to quantitatively investigate inclusion growth dynamics.

The obligate intracellular bacterium Chlamydia elicits a great burden on global public health. C.&#160;trachomatis is the leading bacterial cause of ...

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 ...

Biology

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 ...

Kinetic Visualization of Single-Cell Interspecies Bacterial Interactions

Polymicrobial communities are ubiquitous in nature, yet studying their interactions at the single-cell level is difficult. Thus, a microscopy-based method has been developed for observing interspecies interactions between two bacterial pathogens. The use of this method to interrogate interactions between a motile Gram-negative pathogen, Pseudomonas aeruginosa and a non-motile Gram-positive pathogen, Staphylococcus aureus is demonstrated here. This protocol consists of co-inoculating each species between a coverslip and an agarose pad, which maintains the cells in a single plane and allows for visualization of bacterial behaviors in both space and time.
Furthermore, the time-lapse microscopy demonstrated here is ideal for visualizing the early interactions that take place between two or more bacterial species, including changes in bacterial species motility in monoculture and in coculture with other species. Due to the nature of the limited sample space in the microscopy setup, this protocol is less applicable for studying later interactions between bacterial species once cell populations are too high. However, there are several different applications of the protocol which include the use of staining for imaging live and dead bacterial cells, quantification of gene or protein expression through fluorescent reporters, and tracking bacterial cell movement in both single species and multispecies experiments.

This live-bacterial cell imaging protocol allows for visualization of interactions between multiple bacterial species at the single-cell level over time. Time-lapse imaging allows for observation of each bacterial species in monoculture or coculture to interrogate interspecies interactions in multispecies bacterial communities, including individual cell motility and viability.

Polymicrobial communities are ubiquitous in nature, yet studying their interactions at the single-cell level is difficult. Thus, a microscopy-based method ...

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing molecules of interest to fluorescent proteins such as GFP to study their expression, localization and function. However, GFP and its derivatives are significantly larger and less photostable than organic fluorophores generally used for in vitro experiments, and this can limit the scope of investigation. 
We recently introduced a straightforward, versatile and high-throughput method based on electroporation, allowing the internalization of biomolecules labeled with organic fluorophores into living microorganisms. Here we describe how to use electroporation to internalize labeled DNA fragments or proteins into Escherichia coli and Saccharomyces cerevisi&#230;, how to quantify the number of internalized molecules using fluorescence microscopy, and how to quantify the viability of electroporated cells. Data can be acquired at the single-cell or single-molecule level using fluorescence or FRET. The possibility of internalizing non-labeled molecules that trigger a physiological observable response in vivo is also presented. Finally, strategies of optimization of the protocol for specific biological systems are discussed.

Studies of biomolecules in vivo are crucial for understanding molecular function in a biological context. Here we describe a novel method allowing the internalization of fluorescent biomolecules, such as DNA or proteins, into living microorganisms. Analysis of in vivo data recorded by fluorescence microscopy is also presented and discussed.

The ability to study biomolecules in vivo is crucial for understanding their function in a biological context. One powerful approach involves fusing ...

Live Cell Fluorescence Microscopy to Observe Essential Processes During Microbial Cell Growth

Core cellular processes such as DNA replication and segregation, protein synthesis, cell wall biosynthesis, and cell division rely on the function of proteins which are essential for bacterial survival. A series of target-specific dyes can be used as probes to better understand these processes. Staining with lipophilic dyes enables the observation of membrane structure, visualization of lipid microdomains, and detection of membrane blebs. Use of fluorescent-d-amino acids (FDAAs) to probe the sites of peptidoglycan biosynthesis can indicate potential defects in cell wall biogenesis or cell growth patterning. Finally, nucleic acid stains can indicate possible defects in DNA replication or chromosome segregation. Cyanine DNA stains label living cells and are suitable for time-lapse microscopy enabling real-time observations of nucleoid morphology during cell growth. Protocols for cell labeling can be applied to protein depletion mutants to identify defects in membrane structure, cell wall biogenesis, or chromosome segregation. Furthermore, time-lapse microscopy can be used to monitor morphological changes as an essential protein is removed and can provide additional insights into protein function. For example, the depletion of essential cell division proteins results in filamentation or branching, whereas the depletion of cell growth proteins may cause cells to become shorter or rounder. Here, protocols for cell growth, target-specific labeling, and time-lapse microscopy are provided for the bacterial plant pathogen Agrobacterium tumefaciens. Together, target-specific dyes and time-lapse microscopy enable characterization of essential processes in A. tumefaciens. Finally, the protocols provided can be readily modified to probe essential processes in other bacteria.

Understanding the function of essential processes in bacteria is challenging. Fluorescence microscopy with target-specific dyes can provide key insights into microbial cell growth and cell cycle progression. Here, Agrobacterium tumefaciens is used as a model bacterium to highlight methods for live cell imaging for characterization of essential processes.

Core cellular processes such as DNA replication and segregation, protein synthesis, cell wall biosynthesis, and cell division rely on the function of ...

Multi-color Localization Microscopy of Single Membrane Proteins in Organelles of Live Mammalian Cells

Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a super-resolution technique that allows for the determination of the microcompartments that are accessible for proteins by generating localization and tracking maps of these proteins. Moreover, by multi-color localization microscopy, the localization and tracking profiles of proteins in different subcompartments are obtained simultaneously. The technique is specific for live cells and is based on the repetitive imaging of single mobile membrane proteins. Proteins of interest are genetically fused with specific, so-called self-labeling tags. These tags are enzymes that react with a substrate in a covalent manner. Conjugated to these substrates are fluorescent dyes. Reaction of the enzyme-tagged proteins with the fluorescence labeled substrates results in labeled proteins. Here, Tetramethylrhodamine (TMR) and Silicon Rhodamine (SiR) are used as fluorescent dyes attached to the substrates of the enzymes. By using substrate concentrations in the pM to nM range, sub-stoichiometric labeling is achieved that results in distinct signals. These signals are localized with ~15&#8211;27 nm precision. The technique allows for multi-color imaging of single molecules, whereby the number of colors is limited by the available membrane-permeable dyes and the repertoire of self-labeling enzymes. We show the feasibility of the technique by determining the localization of the quality control enzyme (Pten)-induced kinase 1 (PINK1) in different mitochondrial compartments during its processing in relation to other membrane proteins. The test for true physical interactions between differently labeled single proteins by single molecule FRET or co-tracking is restricted, though, because the low labeling degrees decrease the probability for having two adjacent proteins labeled at the same time. While the technique is strong for imaging proteins in membrane compartments, in most cases it is not appropriate to determine the localization of highly mobile soluble proteins.

Here, we present a protocol for multi-color localization of single membrane proteins in organelles of live cells. To attach fluorophores, self-labeling proteins are used. Proteins, located in different membranes compartments of the same organelle, can be localized with a precision of ~18 nm.

Live-Cell Fluorescence Microscopy to Investigate Subcellular Protein Localization and Cell Morphology Changes in Bacteria

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Monitoring Spatial Segregation in Surface Colonizing Microbial Populations

Live Cell Imaging of Bacillus subtilis and Streptococcus pneumoniae using Automated Time-lapse Microscopy

Single-cell Analysis of Bacillus subtilis Biofilms Using Fluorescence Microscopy and Flow Cytometry

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Kinetic Visualization of Single-Cell Interspecies Bacterial Interactions

Internalization and Observation of Fluorescent Biomolecules in Living Microorganisms via Electroporation

Live Cell Fluorescence Microscopy to Observe Essential Processes During Microbial Cell Growth

Multi-color Localization Microscopy of Single Membrane Proteins in Organelles of Live Mammalian Cells