We thank Professor Hiraoka for sending us strains. We acknowledge support from the Goran Gustafssons Foundation and the Swedish Cancer Society (2008/939).

1. Fission yeast culture
<ol>
	<li>Prepare Edinburgh Minimal Media (EMM) and EMM without ammoniumchloride (EMM-N) 8. To reduce autofluorescence the glucose solution should not be autoclaved but instead filter sterilised using a 0.2 &mu;m filter and subsequently added to the autoclaved media.</li>
	<li>Inoculate fission yeast cells freshly grown on an agar plate with rich media, YEA 8, in 3 mL of EMM liquid media with filter sterilised glucose. Use 13mL tubes with a lightly pushed on cap to ensure good ventilation of the cells. Let the cells grow by shaking them with 225 rpm in 30 &deg;C. Keep the cells growing in log phase (1 X 106 - 2 X 107 cells / mL) for 2 days by counting them using a Burker chamber followed by appropriate dilution, every morning and evening.</li>
	<li>On the day of the experiment make sure to have cells in early log phase, 5 X 106 - 1 X 107 cells / mL.</li>
	<li>To starve the cells for nitrogen during 20 minutes switch from EMM media to EMM-N media. This is done by harvesting 3 ml of cells in a 1.5 mL Eppendorf tube in a bench top centrifuge at maximum 1.5 rcf as centrifugation at a faster speed might induce a stress response in fission yeast 9. Use the double spinning technique, meaning; first spin for 2 min, then turn the Eppendorf tube 180&deg; and spin again for 1.5 rcf 2 min 10. This helps to collect all the cells in one pellet at the bottom of the tube. Wash once with EMM-N and then dissolve the pellet in EMM-N and incubate the cells for 15 minutes at 30 &deg;C shaking at 225 rpm and then continue to point 2. Sample preparation.</li>
</ol>
2. Sample preparation
<ol>
	<li>Harvest 1.5 mL of cells in a 1.5 mL Eppendorf tube in a bench top centrifuge at maximum 1.5 rcf as centrifugation at a faster speed might induce a stress response in fission yeast 9. Use the double spinning technique, meaning; first spin for 2 min, then turn the Eppendorf tube 180&deg; and spin again for 1.5 rcf for 2 min 10. This helps to collect all the cells in one pellet at the bottom of the tube.</li>
	<li>Remove the supernatant, but leave 10-15 &mu;L and resuspend the cells in the remaining media. Alternatively add 10 &mu;L of fresh media to resuspend the cell pellet.</li>
	<li>Make sure to have clear glass slides and cover glass (No 1.5). Normally you do not have to clean them, but make sure they are not full of dust.</li>
	<li>Take out from the freezer an aliquot of a stock solution of 1mg/mL lectin that has been filter sterilised. The lectin solution can be refrozen a few times. The lectin is used to fix the yeast cells at the cover glass.</li>
	<li>Place 5 &mu;L of EMM with filter sterilised glucose on the objective glass.</li>
	<li>Place 5 &mu;L of lectin solution in the corner of the cover glass. Subsequently place 5 &mu;L of cell culture in the same corner, i.e. in the lectin drop. Mix by pipetting a few times and then spread the cell culture-lectin mix throughout the cover glass by using the long side of the pipette tip. Depending on the density of your cell mixture leave everything or suck up excess liquid in the opposite corner of the cover glass.</li>
	<li>Place the cover glass, top up, with one side on the objective glass and the other side on the bench. Let the cover glass dry a little for a few minutes. It should absolutely not be completely dry, but it should not be too much liquid on the glass.</li>
	<li>Place the cover glass with an approximate 70&deg; angel from the objective glass by the drop of EMM. Let go of the cover glass so that it will fall top down into the drop of EMM.</li>
	<li>To seal the edges with silicon grease prepare a 2 mL syringe. Cut the wide end of a 200 &mu;l tip and attach it to the syringe. Fill the syringe with approximately 1 mL of silicon grease. A fine line of silicon is applied to the edges of the cover glass by gently pushing the piston of the syringe. Now you have a small growth chamber of S. pombe cells.</li>
</ol>
3. Microscopy
<ol>
	<li>Initialise the fluorescence microscopy by turning on the mercury/xenon lamp, the microscope and the computer. Place the yeast growth chamber in the fluorescence microscope. Use a 63 X objective or a 100 X objective with NA=1.3 or higher. If an oil objective is used, add oil.</li>
	<li>Use the bright field to find the cells and get a sharp picture.</li>
	<li>The settings for the fluorescence microscopy vary depending on the fluorochromes used to label the yeast cells and the microscope. We use a confocal microscope Zeiss LSM 700 Laser Scanning Microscope with Plan-Apochromat 63x oil objective lens (NA=1.4) with the 16-line average plane scan setting. The pinhole should be set to 1-1.1 Airy units, which gives an optical slice of 0.8 mm. We detect GFP in Track 1 using the filter set for Alexa 488 with a beam splitter at 582 nm, thus detecting wavelengths between 488 and 582 nm. In Track 2 we use a filter set optimal for mCherry using a beam splitter at 578 nm, thus detecting wavelengths between 578 and 600 nm. This means that in Track 2 both the mRFP (SPB) and NM (mCherry) will be detected.</li>
	<li>Take as many pictures needed to be able to measure in 60 different cells for each strain. Usually 15 pictures of independent microscopy fields are enough. It is recommended that a new growth chamber with fresh cells is made if your microscopy time exceeds 60 minutes.</li>
</ol>
4. Quantitative measurement of subcellular distances
<ol>
	<li>Open the pictures in the Zeiss Zen Light Edition program. Using the measurements tool measure the distance in &mu;m between the different fluorochromes in all the cells where all the signals are in the same focal plane. Adjust the light intensity and contrast to identify the centre of the fluorescent signal. This simplified protocol uses only two different colours and hence the SPB and NM are in the same channel. The SPB is singled out by its large round structure in the NM. Transfer the distances to a notepad sheet. Measure in at least 60 cells. Other programs such as ImageJ could also be used to measure the distance, but Zeiss Zen Light is preferred due to its higher resolution of the picture and the ease to zoom in and out using that program. Pictures in lsm format opened in ImageJ will have the two channels on top of each other. To make a picture with both channels, first split the two channels and thereafter merge them. Thereafter the line tool can be used to measure the distances as described above.</li>
	<li>Compare the mean or median subcellular distances between different strains and treatments using a statistical program, for example t-test or Mann-Whitney Rank Sum Test, for example by using the SigmaStat-3.5 software. Frequently the data is not normally distributed since there will be a selection for cells with shorter distances between the fluorescent signals since all the signals need to be in the same focal plane to be measured. A t-test can only be used when the data has a normal distribution while a Mann-Whitney Rank Sum Test allows the comparison of data sets that lack normal distribution. In a t-test the mean of two different datasets are compared while in a Mann-Whitney Rank Sum Test the median is compared.</li>
</ol>
5. Representative Results
Strain PJ1185:(h+ his7+::dis1placR-GFP Chr1[::ura4+ hphMX6 lacO] sid4-mRFP::kanMX6 cut11-mCherry::natMX6 ura4-D18 leu1-32 ade6-DN/N ) was grown in EMM. A sample was withdrawn and mounted in a small growth chamber and pictures were taken (Fig. 1A +N). Subsequently the growth media was replaced with EMM w/o ammoniumchloride (EMM-N), and cells were grown for 15 minutes while shaking. The nitrogen starved cells were then mounted in a growth chamber with EMM-N and pictures were taken (Fig. 1A -N). The measuring tool was used to measure the distance between the locus (GFP) and the SPB (Fig. 1B and Table 1). In addition, the distance between the locus (GFP) and the NM was measured (Fig. 1B and Table 2). The median subcellular distances before and after nitrogen depletion were compared using the SigmaStat-3.5 software (Table 3). There was a statistically significant shift in the localisation of the gene cluster moving away from the NM towards the SBP. The data measuring the distance between the GFP and the SBP had a normal distribution and hence the mean distances (1.777 &mu;m +N and 1,587 &mu;m -N) could be compared using a t-test (Table 1 and 3). There was a significant difference between the two mean values (P = 0.008, t-test). The data measuring the distance between the GFP and NM did not have a normal distribution and hence the median distances (0 &mu;m +N and 0.390 &mu;m -N) were compared using a Mann-Whitney Rank Sum test (Table 2 and 3). There was a significant difference between the two median values (P &lt; 0.001, Mann-Whitney Rank Sum test).
<img alt="Figure 1" src="/files/ftp_upload/3454/3454fig1.jpg" /> 
Figure 1. The localisation of a cluster of genes named Chr1, marked by GFP, changed after the nitrogen starvation. (A) Left column, +N, a representative cell nucleus from a cell grown in EMM right column, -N, a representative cell nucleus from a cell grown in EMM-N. Green is the GFP signal labelling the Chr1 cluster, red is the mRFP and mCherry labelling the SPB and NM, respectively. (B) Same cell nucleus as (A) but now with measured subnuclear distances; yellow: the diameter of the cell nucleus, blue: the distance between SPB and GFP signal and pink: the distance between the GFP signal and the nuclear membrane.
<img alt="Table 1" src="/files/ftp_upload/3454/3454table1a.jpg" /> 
<img alt="Table 1" src="/files/ftp_upload/3454/3454table1b.jpg" /> 
<img alt="Table 1" src="/files/ftp_upload/3454/3454table1c.jpg" /> 
Table 1. The measured subnuclear distances in &mu;m of the PJ1185 strain grown in EMM. First row, diameter of the cell (d), second row, distance between the GFP and the SPB, third row, distance between the GFP and the NM.
<img alt="Table 2" src="/files/ftp_upload/3454/3454table2a.jpg" /> 
<img alt="Table 2" src="/files/ftp_upload/3454/3454table2b.jpg" /> 
<img alt="Table 2" src="/files/ftp_upload/3454/3454table2c.jpg" /> 
Table 2. The measured subnuclear distances in &mu;m of the PJ1185 strain grown in EMM-N. First row, diameter of the cell (d), second row, distance between the GFP and the SPB, third row, distance between the GFP and the NM.
<img alt="Table 3" src="/files/ftp_upload/3454/3454table3.jpg" /> 
Table 3. Descriptive statistics of the observed subnuclear distances in strain PJ1185 before (+N) and after (-N) nitrogen starvation. First row: number of cell measured, second row: mean diameter (d), third row: median diameter (d), forth row: mean distance between the GFP and SPB, fifth row: median distances between the GFP and NM.

<ol>
	<li>Chalfie, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=fluorescent+protein+as+a+marker+for+gene+expression.">fluorescent protein as a marker for gene expression.</a> Science. 263, 802-802 (1994).</li><li>Robinett, C. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+localization+of+DNA+sequences+and+visualization+of+large-scale+chromatin+organization+using+lac+operator/repressor+recognition.">In vivo localization of DNA sequences and visualization of large-scale chromatin organization using lac operator/repressor recognition.</a> J. Cell. 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We have nothing to disclose.

During the last decade the use of live cell imaging to monitor cellular events has become increasingly popular. It started with the use of Green Fluorescence Protein from jellyfish Aequorea victoria and now many different fluorochromes are available emitting fluorescence through a wide spectra from cyan (475 nm) to far red (648 nm) 11. One of the major advantages of live cell imaging over immunofluorescence is that the cells are not fixed by formaldehyde or ethanol/acetone treatment before microscopy, hence avoiding possible artefacts from the fixation process. In addition, live cell imaging offers the possibility to follow individual cells and take pictures at frequent intervals during hours or days of incubation, making it possible to get movies of cellular events 12. Mammalian cells have the advantage of having a larger size, with diameters of around 100 &mu;m, as compared to the smaller yeast cell with a diameter of about 3-4 &mu;m. On the other hand, the advantage with yeast is the easily manipulated genome. Homologous recombination occurs very efficiently in yeast and is used to fuse proteins of interest to different fluorochromes 3. Moreover, using targeted integration of lacO sequences and subsequent expression of LacR-GFP protein allows the detection of a specific part of the genome within the cell nucleus 2. Using S. pombe cells in microscopy has additional advantages since they are natural unicellular organisms that grow with a fast generation time in low-cost cell culture conditions. Moreover, S. pombe is an excellent eukaryotic model organism since it has metazoan homologous genes.
One of the major limitations of this technique is autofluorescence in the yeast cell disturbing the detection of the true signal. This problem can be overcome by using minimal growth media with filter sterilised glucose instead of autoclaved. In addition the yeast cells should be grown for 2 days in log phase before mounting them. The protocol presented here offers a relatively simple, but yet quantitative method to determine the subcellular localisation of proteins within the yeast cell. Moreover by taking pictures at different time points we can follow cellular events.

<table border="1">
	<tbody>
		<tr>
			<td>Name of reagent/equipment</td>
			<td>Company</td>
			<td>Catalogue number</td>
			<td>Comment</td>
		</tr>
		<tr>
			<td>Lectin</td>
			<td>Sigma</td>
			<td>L1395</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Silicon grease</td>
			<td>Dow Corning</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>
			Laser scanning microscopeLSM 700</td>
			<td>Zeiss</td>
			<td>LSM 700</td>
			<td>Other confocal microscope could be used.</td>
		</tr>
		<tr>
			<td>SigmaStat3.5</td>
			<td>Statcon</td>
			<td>SigmaStat3.5</td>
			<td>Any software for statistical analysis could be used.</td>
		</tr>
	</tbody>
</table>

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.

Watch this Scientific Journal Video about Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast at JoVE.com

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

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 ...

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20.2K 조회수.  University of Uppsala. 분열 효모, Schizosaccharomyces pombe의 기본적인 세포 프로세스를 연구하는 좋은 모델 시스템입니다. 여기 우리는 분열 효모의 양적 라이브 세포 분석을 수행하는 방법을 설명합니다. 이 특정 실험에서 우리는 세포 핵 내의 염색체의 조직에 초점을하지만, 방법도 cytosolic 요인을 연구하는 데 사용할 수 있습니다.

비디오: Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

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 ...

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes in cell surface area and cell height, and where cells undergo mitosis and migrate. Furthermore, epithelial cells can also regulate morphogenetic movements in adjacent tissues1. A traditional method to study epithelial cells and tissues involve chemical fixation and histological methods to determine cell morphology or localization of particular proteins of interest. These approaches continue to be useful and provide "snapshots" of cell shapes and tissue architecture, however, much remains to be understood about how cells acquire specific shapes, how various proteins move or localize to specific positions, and what paths cells follow toward their final differentiated fate. High resolution live imaging complements traditional methods and also allows more direct investigation into the dynamic cellular processes involved in the formation, maintenance, and morphogenesis of multicellular epithelial sheets. Here we demonstrate experimental methods from the isolation of animal cap tissues from Xenopus laevis embryos to confocal imaging of epithelial cells and simple measurement approaches that together can augment molecular and cellular studies of epithelial morphogenesis.

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Xenopus embryonic epithelia are an ideal model system to study cell behaviors such as polarity development and shape change during epithelial morphogenesis. Traditional histology of fixed samples is increasingly being complemented by live-cell confocal imaging. Here we demonstrate methods to isolate frog tissues and visualize live epithelial cells and their cytoskeleton using live-cell confocal microscopy.

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes ...

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 ...

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 ...

A Quantitative Fitness Analysis Workflow

Quantitative Fitness Analysis (QFA) is an experimental and computational workflow for comparing fitnesses of microbial cultures grown in parallel1,2,3,4. QFA can be applied to focused observations of single cultures but is most useful for genome-wide genetic interaction or drug screens investigating up to thousands of independent cultures. The central experimental method is the inoculation of independent, dilute liquid microbial cultures onto solid agar plates which are incubated and regularly photographed. Photographs from each time-point are analyzed, producing quantitative cell density estimates, which are used to construct growth curves, allowing quantitative fitness measures to be derived. Culture fitnesses can be compared to quantify and rank genetic interaction strengths or drug sensitivities. The effect on culture fitness of any treatments added into substrate agar (e.g. small molecules, antibiotics or nutrients) or applied to plates externally (e.g. UV irradiation, temperature) can be quantified by QFA.
The QFA workflow produces growth rate estimates analogous to those obtained by spectrophotometric measurement of parallel liquid cultures in 96-well or 200-well plate readers. Importantly, QFA has significantly higher throughput compared with such methods. QFA cultures grow on a solid agar surface and are therefore well aerated during growth without the need for stirring or shaking.
QFA throughput is not as high as that of some Synthetic Genetic Array (SGA) screening methods5,6. However, since QFA cultures are heavily diluted before being inoculated onto agar, QFA can capture more complete growth curves, including exponential and saturation phases3. For example, growth curve observations allow culture doubling times to be estimated directly with high precision, as discussed previously1.
Here we present a specific QFA protocol applied to thousands of S. cerevisiae cultures which are automatically handled by robots during inoculation, incubation and imaging. Any of these automated steps can be replaced by an equivalent, manual procedure, with an associated reduction in throughput, and we also present a lower throughput manual protocol. The same QFA software tools can be applied to images captured in either workflow.
We have extensive experience applying QFA to cultures of the budding yeast S. cerevisiae but we expect that QFA will prove equally useful for examining cultures of the fission yeast S. pombe and bacterial cultures.

Quantitative Fitness Analysis (QFA) is a complementary series of experimental and computational methods for estimating microbial culture fitnesses. QFA estimates the effect of genetic mutations, drugs or other applied treatments on microbe growth. Experiments scaling from focussed analysis of single cultures to thousands of parallel cultures can be designed.

Quantitative Fitness Analysis (QFA) is an experimental and computational workflow for comparing fitnesses of microbial cultures grown in parallel1,2,3,4. ...

Quantitative Analysis of Autophagy using Advanced 3D Fluorescence Microscopy

Prostate cancer is the leading form of malignancies among men in the U.S. While surgery carries a significant risk of impotence and incontinence, traditional chemotherapeutic approaches have been largely unsuccessful. Hormone therapy is effective at early stage, but often fails with the eventual development of hormone-refractory tumors. We have been interested in developing therapeutics targeting specific metabolic deficiency of tumor cells. We recently showed that prostate tumor cells specifically lack an enzyme (argininosuccinate synthase, or ASS) involved in the synthesis of the amino acid arginine1. This condition causes the tumor cells to become dependent on exogenous arginine, and they undergo metabolic stress when free arginine is depleted by arginine deiminase (ADI)1,10. Indeed, we have shown that human prostate cancer cells CWR22Rv1 are effectively killed by ADI with caspase-independent apoptosis and aggressive autophagy (or macroautophagy)1,2,3. Autophagy is an evolutionarily-conserved process that allows cells to metabolize unwanted proteins by lysosomal breakdown during nutritional starvation4,5. Although the essential components of this pathway are well-characterized6,7,8,9, many aspects of the molecular mechanism are still unclear - in particular, what is the role of autophagy in the death-response of prostate cancer cells after ADI treatment? In order to address this question, we required an experimental method to measure the level and extent of autophagic response in cells - and since there are no known molecular markers that can accurately track this process, we chose to develop an imaging-based approach, using quantitative 3D fluorescence microscopy11,12.
Using CWR22Rv1 cells specifically-labeled with fluorescent probes for autophagosomes and lysosomes, we show that 3D image stacks acquired with either widefield deconvolution microscopy (and later, with super-resolution, structured-illumination microscopy) can clearly capture the early stages of autophagy induction. With commercially available digital image analysis applications, we can readily obtain statistical information about autophagosome and lysosome number, size, distribution, and degree of colocalization from any imaged cell. This information allows us to precisely track the progress of autophagy in living cells and enables our continued investigation into the role of autophagy in cancer chemotherapy.

Autophagy is a ubiquitous process that enables cells to degrade and recycle proteins and organelles. We apply advanced fluorescence microscopy to visualize and quantify the small, but essential, physical changes associated with the induction of autophagy, including the formation and distribution of autophagosomes and lysosomes, and their fusion into autolysosomes.

Prostate cancer is the leading form of malignancies among men in the U.S. While surgery carries a significant risk of impotence and incontinence, ...

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 ...

Microscopy of Fission Yeast Sexual Lifecycle

The fission yeast Schizosaccharomyces pombe has been an invaluable model system in studying the regulation of the mitotic cell cycle progression, the mechanics of cell division and cell polarity. Furthermore, classical experiments on its sexual reproduction have yielded results pivotal to current understanding of DNA recombination and meiosis. More recent analysis of fission yeast mating has raised interesting questions on extrinsic stimuli response mechanisms, polarized cell growth and cell-cell fusion. To study these topics in detail we have developed a simple protocol for microscopy of the entire sexual lifecycle. The method described here is easily adjusted to study specific mating stages. Briefly, after being grown to exponential phase in a nitrogen-rich medium, cell cultures are shifted to a nitrogen-deprived medium for periods of time suited to the stage of the sexual lifecycle that will be explored. Cells are then mounted on custom, easily built agarose pad chambers for imaging. This approach allows cells to be monitored from the onset of mating to the final formation of spores.

We provide a reproducible basic method for the long-term microscopy of the fission yeast sexual lifecycle. With minor adjustments described, the presented protocol allows research focus on different steps of the reproductive process.

The fission yeast Schizosaccharomyces pombe has been an invaluable model system in studying the regulation of the mitotic cell cycle progression, the ...

Spatiotemporal Analysis of Cytokinetic Events in Fission Yeast

Cytokinesis, the final step in cell division is critical for maintaining genome integrity. Proper cytokinesis is important for cell differentiation and development. Cytokinesis involves a series of events that are well coordinated in time and space. Cytokinesis involves the formation of an actomyosin ring at the division site, followed by ring constriction, membrane furrow formation and extra cellular matrix remodeling. The fission yeast, Schizosaccharomyces pombe (S.&#160;pombe)&#160;is a well-studied model system that has revealed with substantial clarity the initial events in cytokinesis. However, we do not understand clearly how different cytokinetic events are coordinated spatiotemporally. To determine this, one needs to analyze the different cytokinetic events in great details in both time and in space. Here we describe a microscopy approach to examine different cytokinetic events in live cells. With this approach it is possible to time different cytokinetic events and determine the time of recruitment of different proteins during cytokinesis. In addition, we describe protocols to compare protein localization, and distribution at the site of cell division. This is a basic protocol to study cytokinesis in fission yeast and can also be used for other yeasts and fungal systems.

The fission yeast, Schizosaccharomyces pombe&#160;is an excellent model system to study cytokinesis, the final stage in cell division. Here we describe a microscopy approach to analyze different cytokinetic events in live fission yeast cells.

Cytokinesis, the final step in cell division is critical for maintaining genome integrity. Proper cytokinesis is important for cell differentiation and ...

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 ...