We would like to thank Jonathan Cooper Lab at Fred Hutchinson Cancer Research Center for the permission to use their cryostat. This work was supported by NIH and the W.M. Keck Foundation. BM is a Gordon and Betty Moore fellow of Life Science Research Foundation. We are grateful to Daniel Gottschling for sharing his fixing protocol with us.

1. Fixing Yeast Communities
<ol>
	<li>Grow yeast communities on a membrane filter (e.g., Millipore MF membrane filter; Figure 2A). This protocol corresponds to a typical community of less than 2x108 cells on a 6 mm-diameter membrane filter.</li>
	<li>Use a piece of Whatman 541 filter paper to cover the bottom of a 1 cm-diameter well (Figure 2B). This Whatman paper will be used as a carrier to transfer the community in later stages. Add 1 ml 100% methanol to the well, and leave the well in -20 &deg;C for the temperature to equilibrate.</li>
	<li>Put liquid N2 in a dewar. Freeze community in liquid nitrogen by dipping the community for at least 15 sec in liquid N2 using a tweezer (Figure 2C).</li>
	<li>Transfer frozen community to the well of -20 &deg;C methanol (Figure 2D). Ensure that the community is kept horizontal when submerging it in the methanol. Wait 20 min. The MF membrane filter starts to disintegrate in methanol.</li>
	<li>Transfer community using the carrier Whatman 541 filter paper from the methanol well to a cold Petri dish kept at -20 &deg;C for methanol evaporation (Figure 2E). When transferring, keep the community on top of carrier Whatman filter and draw away extra drops of methanol using another piece of thick Whatman paper before putting in the cold Petri dish. This will ensure that evaporation time is consistent from sample to sample.</li>
	<li>Wait 4-6 hr for evaporation until the carrier Whatman filter looks dry. Insufficient evaporation leaves residual methanol which interferes with sectioning. Over-drying causes cracks and deformation in communities.</li>
	<li>Take the sample out of -20 &deg;C freezer. Cut away Whatman 541 filter paper not covered by community.</li>
	<li>Make a rectangular cubic aluminum-foil container which is large enough such that the sample has at least ~2 mm margin from each side (Figure 2F). Place the community on the bottom surface of the aluminum-foil container; immediately cover the sample with OCT to 3~4 mm above community height. If important, the orientation of the sample can be adjusted in this step with regard to the sides of the container. Wait 10 min.</li>
	<li>Place the OCT-embedded sample on a metallic plate on dry ice to freeze. Store the frozen communities at -20 &deg;C or -80 &deg;C freezer.</li>
</ol>
2. Sectioning Frozen Communities for Fluorescence Imaging
<ol>
	<li>Set the temperature of cryosection chamber to -25 &deg;C and leave the samples in the chamber until its temperature reaches the temperature of the chamber.</li>
	<li>Mount the sample onto a cryo-mount after applying a drop of OCT on the mount. Spread OCT uniformly on the side of sample with exposed Whatman filter paper (bottom of the container) so that OCT thickness is 2-3 mm. Let OCT freeze inside the cryostat chamber before sectioning. Ensuring an OCT margin of 2-3 mm on all sides of the sample helps to preserve the integrity of sections.</li>
	<li>The sides of the OCT sample block can be used to align the sample. If the sample is not aligned with respect to the blade, sectioning will start from a corner or side of the sample. Adjust the mount to ensure that the blade cuts a perfectly vertical (or horizontal) face of the sample block.</li>
	<li>Typical sections for yeast communities are ~14 &mu;m-thick. After cutting a section, use a microscope slide kept at room temperature to collect the section, by slowly approaching the slide to the cut section. The section will melt and transfer to the microscope slide. Up to 18 sections can be transferred onto one slide.</li>
	<li>Store the slides at cold temperatures, preferably -20 &deg;C, before imaging. Loss of signal is minimal when kept for one week at 4 &deg;C. For maximum fluorescence signal, imaging is preferably done immediately after sectioning.</li>
</ol>

<ol>
	<li>Tolker-Nielsen, T., Molin, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatial+Organization+of+Microbial+Biofilm+Communities.">Spatial Organization of Microbial Biofilm Communities.</a> Microbial Ecology. 40 (2), 75-84 (2000).</li><li>Moller, 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=+In+Situ+Gene+Expression+in+Mixed-Culture+Biofilms:+Evidence+of+Metabolic+Interactions+between+Community+Members."> In Situ Gene Expression in Mixed-Culture Biofilms: Evidence of Metabolic Interactions between Community Members.</a> Appl. Environ. Microbiol. 64 (2), 721-732 (1998).</li><li>Lepp, P. W., 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=Methanogenic+Archaea+and+human+periodontal+disease.">Methanogenic Archaea and human periodontal disease.</a> Proceedings of the National Academy of Sciences of the United States of America. 101 (16), 6176-6181 (2004).</li><li>V&#225;chov&#225;, L., 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=Architecture+of+developing+multicellular+yeast+colony:+spatio-temporal+expression+of+Ato1p+ammonium+exporter.">Architecture of developing multicellular yeast colony: spatio-temporal expression of Ato1p ammonium exporter.</a> Environmental Microbiology. 11 (7), 1866-1877 (2009).</li><li>Min&#225;rikov&#225;, L., 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=Differentiated+Gene+Expression+in+Cells+within+Yeast+Colonies.">Differentiated Gene Expression in Cells within Yeast Colonies.</a> Experimental Cell Research. 271 (2), 296-304 (2001).</li><li>Shou, W., 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=Synthetic+cooperation+in+engineered+yeast+populations.">Synthetic cooperation in engineered yeast populations.</a> Proc. Natl. Acad. Sci. U.S.A. 104, 1877-1882 (2007).</li><li>Momeni, B., 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=Cooperation+generates+a+unique+spatial+signature+in+microbial+communities.">Cooperation generates a unique spatial signature in microbial communities.</a> , (2012).</li><li>Kiernan, J., ed, <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Histological and Histochemical Methods: Theory and Practice. , 4 (2008).</li><li>Piccirillo, 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=The+Rim101p/PacC+Pathway+and+Alkaline+pH+Regulate+Pattern+Formation+in+Yeast+Colonies.">The Rim101p/PacC Pathway and Alkaline pH Regulate Pattern Formation in Yeast Colonies.</a> Genetics. 184 (3), 707-716 (2010).</li></ol>

No conflicts of interest declared.

The procedure presented here offers a way to inspect the internal structure of yeast communities. The methanol fixing step makes the yeast colony rigid8 and preserves the integrity of the colony; however, it also causes shrinkage8 that should be taken into account in interpreting the results. We typically see around 30% shrinkage in height of yeast colonies, compared to colonies directly embedded in OCT without fixing9.
To avoid shrinkage, an alternativ...

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>100% Methanol</td>
 <td>J.T. Baker</td>
 <td>9070-01</td>
 <td></td>
 </tr>
 <tr>
 <td>Tissue-Tek O.C.T. Compound</td>
 <td>Andwin Scientific</td>
 <td>4583</td>
 <td></td>
 </tr>
 <tr>
 <td>Whatman Grade No. 541 Quantitative Filter Paper</td>
 <td>Whatman</td>
 <td>1541-047 </td>
 <td></td>
 </tr>
 <tr>
 <td>Millipore-MF Membrane Filter</td>
 <td>Millipore</td>
 <td>HAWP04700</td>
 <td></td>
 </tr>
 <tr>
 <td>Cryostat</td>
 <td>Leica</td>
 <td>CM 1510 S</td>
 <td></td>
 </tr>
 </tbody>
</table>

cryosectioning yeast communities for examining fluorescence patterns

Microbes typically live in communities. The spatial organization of cells within a community is believed to impact the survival and function of the community1. Optical sectioning techniques, including confocal and two-photon microscopy, have proven useful for observing spatial organization of bacterial and archaeal communities2,3. A combination of confocal imaging and physical sectioning of yeast colonies has revealed internal organization of cells4. However, direct optical sectioning using confocal or two-photon microscopy has been only able to reach a few cell layers deep into yeast colonies. This limitation is likely because of strong scattering of light from yeast cells4.
Here, we present a method based on fixing and cryosectioning to obtain spatial distribution of fluorescent cells within Saccharomyces cerevisiae communities. We use methanol as the fixative agent to preserve the spatial distribution of cells. Fixed communities are infiltrated with OCT compound, frozen, and cryosectioned in a cryostat. Fluorescence imaging of the sections reveals the internal organization of fluorescent cells within the community.
Examples of yeast communities consisting of strains expressing red and green fluorescent proteins demonstrate the potentials of the cryosectioning method to reveal the spatial distribution of fluorescent cells as well as that of gene expression within yeast colonies2,3. Even though our focus has been on Saccharomyces cerevisiae communities, the same method can potentially be applied to examine other microbial communities.

Fluorescence images of vertical cross-sections of yeast communities containing red- and green-tagged competitive populations6 are shown in Figure 3. These communities consist of two prototrophic strains of yeast and their competition for shared resources, including space, leads to columnar patterns7, as displayed in their vertical cross-sections. Resolutions down to a single cell can be resolved in cross-sections. Figure 3 compares cross-sections of replicate ...

Watch this Scientific Journal Video about Cryosectioning Yeast Communities for Examining Fluorescence Patterns at JoVE.com

Cryosectioning Yeast Communities for Examining Fluorescence Patterns

We present a protocol for freezing and cryosectioning yeast communities to observe internal patterns of fluorescent cells. The method relies on methanol-fixing and OCT-embedding to preserve the spatial distribution of cells without inactivating fluorescent proteins within a community.

Microbes typically live  in communities. The spatial organization  of cells within a community is believed to impact the survival and function of the  ...

cryosectioning-yeast-communities-for-examining-fluorescence-patterns

Research

JoVE Journal

Biology

8.8K Views.  Fred Hutchinson Cancer Research Center. Vi presentiamo un protocollo per congelare e criosezionamento comunità di lievito per osservare i modelli interni di cellule fluorescenti. Il metodo si basa su di fissaggio e metanolo-OCT-incorporamento per preservare la distribuzione spaziale di celle senza inattivare proteine ​​fluorescenti all'interno di una comunità.

Video: Cryosectioning Yeast Communities for Examining Fluorescence Patterns

Flash Freezing and Cryosectioning E12.5 Mouse Brain

Demonstrated in this video are the techniques for flash freezing and sectioning embryonic brain tissue from mouse. Useful tips for using the cryostat are given, including troubleshooting methods that can be used while cutting to ensure that the resultant tissues sections are free of cracks and other distortions.

Microbial Communities in Nature and Laboratory - Interview

Biology of Microbial Communities  - Interview

A Microfluidic Device with Groove Patterns for Studying Cellular Behavior

We describe a microfluidic device with microgrooved patterns for studying cellular behavior. This microfluidic platform consists of a top fluidic channel and a bottom microgrooved substrate. To fabricate the microgrooved channels, a top poly(dimethylsiloxane) (PDMS) mold containing the impression of the microfluidic channels was aligned and bonded to a microgrooved substrate. Using this device, mouse fibroblast cells were immobilized and patterned within microgrooved substrates (25, 50, 75, and 100 &mu;m wide). To study apoptosis in a microfluidic device, media containing hydrogen peroxide, Annexin V, and propidium iodide was perfused into the fluidic channel for 2 hours. We found that cells exposed to the oxidative stress became apoptotic. These apoptotic cells were confirmed by Annexin V that bound to phosphatidylserine at the outer leaflet of the plasma membrane during the apoptosis process. Using this microfluidic device with microgrooved patterns, the apoptosis process was observed in real-time and analyzed by using an inverted microscope containing an incubation chamber (37&deg;C, 5% CO2). Therefore, this microfluidic device incorporated with microgrooved substrates could be useful for studying the cellular behavior and performing high-throughput drug screening.

We describe a protocol for the fabrication of microfluidic devices that can enable cell capture and culture. In this approach patterned microstructures such as grooves within microfluidic channels are used to create low shear stress regions within which cell can dock.

We describe a microfluidic device with microgrooved patterns for studying cellular behavior.  This microfluidic platform consists of a top fluidic channel ...

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 ...

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

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

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

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

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

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

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 ...

Acquiring Fluorescence Time-lapse Movies of Budding Yeast and Analyzing Single-cell Dynamics using GRAFTS

Fluorescence time-lapse microscopy has become a powerful tool in the study of many biological processes at the single-cell level. In particular, movies depicting the temporal dependence of gene expression provide insight into the dynamics of its regulation; however, there are many technical challenges to obtaining and analyzing fluorescence movies of single cells. We describe here a simple protocol using a commercially available microfluidic culture device to generate such data, and a MATLAB-based, graphical user interface (GUI) -based software package to quantify the fluorescence images. The software segments and tracks cells, enables the user to visually curate errors in the data, and automatically assigns lineage and division times. The GUI further analyzes the time series to produce whole cell traces as well as their first and second time derivatives. While the software was designed for S. cerevisiae, its modularity and versatility should allow it to serve as a platform for studying other cell types with few modifications.

We present a simple protocol to obtain fluorescence microscopy movies of growing yeast cells, and a GUI-based software package to extract single-cell time series data. The analysis includes automated lineage and division time assignment integrated with visual inspection and manual curation of tracked data.

Fluorescence time-lapse microscopy has become a powerful tool in the study of many biological processes at the single-cell level. In particular, movies ...

Cryosectioning Method for Microdissection of Murine Colonic Mucosa

The colonic mucosal tissue provides a vital barrier to luminal antigens. This barrier is composed of a monolayer of simple columnar epithelial cells. The colonic epithelium is dynamically turned over and epithelial cells are generated in the stem cell containing crypts of Lieberk&#252;hn. Progenitor cells produced in the crypt-bases migrate toward the luminal surface, undergoing a process of cellular differentiation before being shed into the gut lumen. In order to study these processes at the molecular level, we have developed a simple method for the microdissection of two spatially distinct regions of the colonic mucosa; the proliferative crypt zone, and the differentiated surface epithelial cells. Our objective is to isolate specific crypt and surface epithelial cell populations from mouse colonic mucosa for the isolation of RNA and protein.

Here we describe a simple method for the isolation of spatially distinct murine colonic epithelial surface cells from the underlying crypt-base cells.

The colonic mucosal tissue provides a vital barrier to luminal antigens. This barrier is composed of a monolayer of simple columnar epithelial cells. The ...