We thank Michael VanNieuwenhze (Indiana University) for the gift of the FDAAs used in Figure 2 and Figure 4. We thank members of the Brown lab for feedback during the preparation of this manuscript. Research in the Brown lab on A. tumefaciens cell growth and division is supported by the National Science Foundation (IOS1557806).

1. Growth of A. tumefaciens Strains
<ol>
	<li>Culturing A. tumefaciens strains
	<ol>
		<li>Use a sterile wooden stick or pipet tip to inoculate 1 mL of ATGN growth media (see materials list for the recipe) with a single colony of the desired strain. 
		NOTE: For A. tumefaciens depletion strains, the ATGN should contain 1 mM IPTG as an inducer to maintain biosynthesis of the essential protein.</li>
		<li>Grow the A. tumefaciens strains overnight in ATGN at 28 &#176;C with shaking at 225 rpm.</li>
		<li>Measure the optical density of the cells at 600 nm (OD600) using a spectrophotometer. Dilute the cell culture to an OD600 = ~0.2 and continue to grow until OD600 = ~0.6 is reached. For depletion strains, continue to grow with inducer until OD600 = ~0.6 is reached.</li>
		<li>Use wildtype and mutant strains of A. tumefaciens at OD600 = ~0.6 for target-specific staining and/or time-lapse microscopy (see sections 2 and 3.1). For depletion strains, wash out the inducer (see section 1.2.)</li>
	</ol>
	</li>
	<li>Removal of inducer for characterization of A. tumefaciens protein depletion strains
	<ol>
		<li>Pellet 1 mL of exponential culture (OD600 = ~0.4-0.6) by centrifugation at 7,000 x g for 5 min in a desktop centrifuge at room temperature.</li>
		<li>To wash, remove supernatant and resuspend the pellet in fresh media without inducer and pellet the cells as described above (1.2.1). Wash the cells a total of 3 times in fresh media.</li>
		<li>Resuspend the final pellet in fresh media and concentrate the cells to an OD600 = ~0.8 for immediate staining with target specific dyes (section 2 and Figure 4B-D) or time-lapse microscopy (section 3.2 and Figure 4A). Alternatively, pre-deplete the cells for the desired amount of time by growing the cells in media without inducer prior to staining and imaging of the cells.</li>
	</ol>
	</li>
</ol>
2. Target-specific Staining of A. tumefaciens Cells
<ol>
	<li>Fluorescent d-amino acid cell wall labeling 
	NOTE: FDAAs are non-toxic fluorescent d-amino acids which are readily incorporated into the A. tumefaciens peptidoglycan. This simple labeling procedure probes growth patterning in live cells6. Protocols for synthesis of four FDAAs are available38.
	<ol>
		<li>Pellet 500 &#181;L of exponential culture (OD600 = ~0.4-0.6) by centrifugation at 7,000 x g for 5 min in a desktop centrifuge. Resuspend the cell pellet in 100 &#181;L of fresh media.</li>
		<li>Add 5 &#181;L of 5 mM FDAA to washed and concentrated cells and incubate for 2 min in the dark. 
		NOTE: Limit FDAA exposure to light. The time of incubation will vary depending on the growth rate of the strain. Typically, incubation times should be 5-10% of the doubling time6.</li>
		<li>Pellet the cells by centrifugation and wash cell pellets with phosphate buffered saline (PBS) three times.</li>
		<li>Resuspend pellet in ~50 &#181;L PBS. 
		NOTE: The volume of PBS used for resuspension may vary based on pellet size.</li>
		<li>Apply 0.8-1 &#181;L of cells to an agarose pad and image using epifluorescence microscopy immediately (See sections 3.1 and 3.2).
		<ol>
			<li>Alternatively, stop further label incorporation by fixing the cells with ice-cold 70% ethanol. Resuspend the cell pellet in 1 mL of ice-cold 70% ethanol and incubate on ice for 15 minutes.</li>
			<li>Collect cells by centrifugation and resuspend in a small volume of PBS for imaging at a later time. Store cell suspensions at 4 &#176;C and image within 48 hours.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>DNA staining with DAPI or Orange stain 
	NOTE: To observe DNA structure, cells are labeled with either DAPI or Orange (dead cell stain). Although classically described as a &#34;dead-cell stain&#34;, Orange is permeant, photostable, and does not affect the growth of bacterial cells, enabling live-cell DNA imaging10. In A. tumefaciens, Orange labeling works well in living cells and is suitable for time-lapse microscopy (Figure 3). In contrast, the ultraviolet (UV) light exposure necessary for imaging DAPI-stained cells is phototoxic (Figure 3) and thus DAPI staining is best suited for staining live cells for immediate imaging or staining fixed cells.
	<ol>
		<li>DAPI staining of cells
		<ol>
			<li>Pellet 1 mL of exponential culture (OD600 = ~0.4-0.6) by centrifugation at 7,000 x g for 5 min in a desktop centrifuge.</li>
			<li>Optionally, to fix cells prior to staining, resuspend the cell pellet in 1 mL of 70% ice-cold ethanol. Incubate on ice for 10-15 min. Collect the cells by centrifugation as described in 2.2.1.1.</li>
			<li>Resuspend the cell pellet in 1 mL of PBS containing 1 &#181;L of DAPI stock solution (1 mg/mL; see Materials Table). Mix gently by pipetting and incubate for 5 minutes in the dark.</li>
			<li>Pellet cells by centrifugation at 7,000 x g for 5 min and resuspend in 1 mL of PBS to remove excess DAPI. Wash the cells two more times and resuspend the pellet in 50 &#181;L PBS or media.</li>
			<li>Apply 0.8-1 &#181;L of cells to an agarose pad and image using epifluorescence microscopy immediately. See sections 3.1 and 3.2. 
			NOTE: This protocol is not optimal for time-lapse microscopy to observe chromosome dynamics (Figure 3). Consider using Orange staining as an alternative (see section 2.2.2).</li>
		</ol>
		</li>
		<li>Orange staining of live cells</li>
		<li>Pellet 1 mL of exponential culture (OD600 = ~0.4-0.6) by centrifugation at 7,000 x g for 5 min in a desktop centrifuge. Resuspend cell pellet in 1 mL of PBS containing 1 &#181;L of Orange stock solution (see materials list). Mix gently by pipetting and incubate for 5 min in the dark.</li>
		<li>Pellet cells by centrifugation and resuspend in 1 mL of PBS to remove excess Orange. Wash the cells two more times and resuspend the pellet in 50 &#181;L PBS.</li>
		<li>Apply 0.8-1 &#181;L of cells to an agarose pad and image using epifluorescence microscopy immediately. See sections 3.1 and 3.2. 
		NOTE: This protocol works well with live cells and is suitable for time-lapse microscopy to observe chromosome dynamics (Figure 3). If it is not convenient to image immediately, cells may be fixed in ice-cold 70% ethanol (see section 2.2.1.2).</li>
	</ol>
	</li>
	<li>Membrane labeling 
	NOTE: The lipophilic styryl fluorescent dye 4-64 has been used extensively to observe the membrane of bacterial cells. In contrast to many bacteria, 4-64 labeled A. tumefaciens cells do not label uniformly, but rather frequently exhibit a &#34;horseshoe&#34; pattern14,15. Remarkably, the growth pole is nearly devoid of stain whereas the old pole is intensely labeled. Thus, 4-64 enables visualization of both cell growth patterns and membrane structure in A. tumefaciens.
	<ol>
		<li>Add FM 4-64 to a final concentration of 8 &#181;g/mL in 1 mL of exponential phase cell culture and incubate at room temperature for 5 minutes in the dark.</li>
		<li>Pellet labeled cells by centrifugation at 7,000 x g for 5 minutes in a desktop centrifuge and resuspend cell pellet in 1 mL of PBS. Wash the cells a total of three times to remove excess dye.</li>
		<li>Resuspend cells in a small volume of PBS and spot on an agarose pad to image immediately. (See sections 3.1 and 3.2) 
		CAUTION: Cells must be imaged immediately to observe characteristic labeling patterns.</li>
	</ol>
	</li>
</ol>
3. Imaging of A. tumefaciens Cells
<ol>
	<li>Agarose pad preparation 
	NOTE: Figure 1 contains an image sequence (panel A) and schematic (panel B) of a typical agarose pad prepared for time-lapse microscopy. Agarose pads are typically prepared as needed.
	<ol>
		<li>3.1.1. Use a coverslip as a guide and cut a 22 mm x 22 mm square of laboratory film (see materials list) by running a scalpel around the edges.</li>
		<li>Cut a square out of the center of the laboratory film leaving a ~2-5 mm border to serve as a gasket for the agarose pad. Discard the center cutout.</li>
		<li>Place the film gasket onto a glass slide (75 mm x 25 mm) cleaned with an ammonia and alcohol-free cleaner (see materials list) and heat until the film is slightly melted onto glass (top image in Figure 1A). To melt laboratory film, use the edge of a heat block set to 70 &#176;C or melt lightly over a flame.</li>
		<li>Prepare agarose solution by mixing ~0.075 g agarose (see materials list) in 5 mL of media in a small flask. Heat the solution in a microwave with periodic swirling to mix until the agarose is dissolved and the solution is clear. Keep the agarose solution at 55-70 &#176;C and use for construction of multiple agarose pads within 48 hours.</li>
		<li>Pipette media containing 1.2-1.5% agarose into the center of the gasket. 
		NOTE: The volume of media is typically 50 - 60 &#181;L but will vary based on gasket size. Adding media to a cold slide can cause the agarose to solidify too quickly, thus the slide should be kept on a warm surface. The edge of a heat block set to 70 &#176;C works well. Water agarose or buffered agarose solutions such as phosphate buffered saline (PBS) can be used instead of media containing agarose for applications in which continued cell growth is not required. For imaging depletion strains, inducer can be present or absent in the media. Presence of inducer can be used as a control whereas the absence of inducer will reveal the depletion phenotype.</li>
		<li>Place a coverslip over the gasket to evenly distribute the agarose.</li>
		<li>Place slide on a cool, level surface to solidify for ~2 min. Avoid overdrying of the agarose pad as this will result in a wrinkled surface on the agarose pad and pooling of the cell suspension when applied to the surface.</li>
		<li>Carefully slide the coverslip off the agarose pad. 
		Caution: Do not rush this step. The agarose pad can easily tear and an uneven agarose pad can make imaging difficult.</li>
		<li>Allow the agarose pad to air dry for 1-2 min at room temperature until the surface of the pad appears dry. Using a scalpel, remove a small strip of agarose to create an air pocket (middle image, Figure 1A); the air pocket is typically ~2 mm x 7-10 mm and A. tumefaciens cells tend to grow best near the air pocket (Figure 1C). 
		NOTE: For imaging of fixed cells or under conditions where growth is not monitored, an air pocket is not necessary.</li>
		<li>Spot 0.8-1 &#181;L of cells on the agarose pad and cover with a new coverslip. Gently place the coverslip over the top of the agarose pad to distribute the cells across the surface of the agarose pad.</li>
		<li>Seal the edges of the coverslip using melted VALAP (see Materials Table for recipe; Figure 1A, bottom image). Failure to seal along all the edges and corners of the coverslip can cause drying of the agarose pad, which will lead to the drifting of cells during imaging. NOTE: Sealing of the coverslip is only needed for long-term time-lapse imaging.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56497/56497fig1.jpg" /> 
Figure 1: Agarose pad preparation. (A) Image sequence of the agarose pad preparation protocol. Image 1 is a slide with the laboratory film gasket. In image 2, the agarose pad and air pocket are visualized. Finally, image 3 shows the complete agarose pad with the cells under a coverglass and sealed with VALAP. (B) A schematic of an agarose pad for time-lapse microscopy is provided. Key features of the agarose pad are labeled on the schematic. (C) The air pocket promoted growth of A. tumefaciens on agarose pads. Images of wildtype A. tumefaciens cells grown on an agarose pad for 20 hours are shown. Images were taken at positions increasingly distant from the air pocket. The distance from the closest edge of the image to the air pocket is shown above each image. Scale bar = 10 &#181;m. <a href="//cloudfront.jove.com/files/ftp_upload/56497/56497fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="2">
	<li>Imaging 
	NOTE: Differential interference contrast, phase contrast, and epifluorescence imaging is performed with an inverted microscope equipped with hardware-based autofocusing, an automated stage, standard filters, an LED light source, 60X oil immersion objectives (1.4 NA) for phase contrast or differential interference contrast (DIC), and a 1K electron-multiplying charge-coupled-device (EMCCD) camera. The microscope should be placed in a temperature controlled room. Alternatively, a stage warmer or chamber can be used to maintain consistent temperatures during imaging.
	<ol>
		<li>Epifluorescence microscopy 
		NOTE: For fluorescence imaging of live cells, it is necessary to limit the number of image acquisitions and optimize the exposure time to minimize photobleaching and phototoxicity. For imaging of each dye, it is recommended to find an optimal exposure time which provides sufficient fluorescence detection but does not lead to photobleaching or phototoxicity. In Figures 2-4, exposure times of 200 ms were used for all fluorescence images. For the time-lapse sequences shown in Figure 3, images were acquired every 10 min for 3 h.
		<ol>
			<li>Place immersion oil on the desired objective and place the inverted slide into the slide holder on the stage. Use the focusing knobs to bring the cells into focus.</li>
			<li>Acquire images in phase (or DIC) and the desired fluorescence filter. 
			NOTE: The maximal excitation and emission wavelengths for the stains used in Figure 2, Figure 3, and Figure 4 are as follows: HADA (405/460), NADA (450/555), TADA (555/570), 4-64 (515/640), DAPI (360/460), Orange (547/570).</li>
		</ol>
		</li>
		<li>Time-lapse microscopy 
		NOTE: During time-lapse microscopy the following features are computer controlled: x, y, and z position, shutters, and fluorescence filters. A hardware-based auto-focus system is optimal to maintain focus during time-lapse imaging. Alternatively, a software-based auto-focusing loop can be used.
		<ol>
			<li>Place immersion oil on the desired objective and place the inverted slide into the slide holder on the stage. Use the focusing knobs to bring the cells into focus.</li>
			<li>Optionally: Acquire multiple (x,y) positions. Randomly select 10 fields of cells in close proximity to the air pocket of the agarose pads.</li>
			<li>Set-up the time sequence acquisition to image in phase or DIC at the desired time interval. 
			NOTE: For the time-lapse sequence shown in Figure 4, DIC images were acquired with 30 ms exposure every 10 min for 14 h. When using fluorescence imaging during time-lapse microscopy, adjust the acquisition interval and exposure time to minimize photobleaching and phototoxicity.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

This protocol contains a series of procedures for the investigation of A. tumefaciens wildtype, mutant, and depletion strains. It is worth noting that all of the procedures listed in the protocol section can be readily adapted for other bacterial strains with additional modifications to account for growth media, temperatures, and growth rates.
The use of target-specific dyes is a valuable tool for providing a detailed characterization of cell-cycle events in bacterial cells. Here, pep...

Progression through the bacterial cell cycle requires the coordination of many processes including membrane and cell wall biosynthesis, DNA replication and segregation, and cell division. To fully understand the complexity of bacterial cell biology, it is necessary to study these essential events; however, this is a non-trivial task since cell viability is compromised when key components of these pathways are mutagenized. Epifluorescence microscopy coupled with target-specific dyes is a powerful approach to probe these essential processes in wildtype and mutant bacterial strains.
Peptidoglycan-specific dyes include fluorescent antibiotics (vancomycin-FL, bocillin-FL) and fluorescent-d-amino acids (for example, 7-hydroxycoumarin-3-carboxylic acid-3-amino-d-alanine, HADA; 4-chloro-7-nitrobenzofurazan-3-amino- d-alanine, NADA; tetramethylrhodamine-3-amino-d-alanine; TADA). In Gram-positive bacteria, the use of sublethal concentrations of fluorescent antibiotic analogs to probe sites of peptidoglycan biosynthesis has been an effective strategy to reveal peptidoglycan insertion patterns1,2,3,4. While fluorescent vancomycin labeling has been used to gain insights into the peptidoglycan insertion patterns in fixed Gram-negative bacteria5, the outer membrane generally provides a permeability barrier that prevents the use of fluorescent antibiotics as a probe for peptidoglycan biosynthesis in live cells. In contrast, short pulses of fluorescent-d-amino acids or d-amino acids with biorthogonal functional groups covalently label regions of recent peptidoglycan insertion in a wide range of living bacterial cells6,7. Patterns of peptidoglycan insertion that have been observed with synthetic d-amino acids include punctate and septal (Escherichia coli and Bacillus subtilis), polar and septal (Agrobacterium tumefaciens and Listeria monocytogenes), septal only (Staphylococcus aureus), and apical (Streptomyces venezuelae)6,7. These observations indicate that bacteria exhibit diverse patterns of cell wall biogenesis and that the use of synthetic d-amino acids as probes for examining growth patterning is a valuable strategy in many bacteria.
Dyes that label bacterial chromosomes include the deoxyribonucleic acid (DNA) specific minor groove binder (4,6-diamidino-2-phenylindole; DAPI) and high affinity cyanine dyes (Green and Orange; see materials list). DAPI staining of fixed cells assists in enumeration of bacteria from environmental samples8, whereas DAPI staining of live cells is used to indicate bacterial viability9. In contrast, cyanine dyes such as Orange and Green are frequently described as membrane impermeant &#34;dead&#34; cell stains to enumerate non-viable cells9. Remarkably, when these reagents are used to probe the morphology of the bacterial nucleoid during cell growth, DAPI, Orange, and Green were all shown to be membrane permeant and capable of labeling live cells10. In live E. coli cells, DAPI staining of DNA appears diffuse due to auto-fluorescence from the cytoplasm and repeated exposures of DAPI-stained cells to ultraviolet (UV) light perturbs the nucleoid structure10. Staining E. coli or B. subtilis with Orange reveals that this dye is membrane permeant and provides long-lasting fluorescence upon binding to DNA in live cells without impacting cell growth, DNA replication, or chromosome segregation10. These observations suggest that cyanine DNA dyes can be used to monitor the morphology of nucleoids during cell growth in many bacteria.
Phospholipid-specific stryl dyes such as N-(3-triethylammoniumpropyl) -4-(6-(4-(diethylamino) phenyl)hexatrienyl)pyridinium dibromide (4-64; see materials list) are cationic compounds and associate preferentially with negatively charged phospholipids such as cardiolipin and phosphatidylglycerol11. Distinct patterns are observed when 4-64 is used to label the membrane of different bacteria. In Escherichia coli, 4-64 is enriched in the poles, in bands along the lateral wall, and in the division sites of late pre-divisional cells12. In Bacillus subtilis, 4-64 labeling enables the visualization of lipid spirals13. In Agrobacterium tumefaciens, 4-64 labels the outer membrane and is observed in a characteristic &#34;horseshoe&#34; pattern in which the growth pole is devoid of labeling14,15. These observations indicate that these bacteria exhibit heterogeneous lipid distributions due to the presence of lipid domains which contribute to cellular asymmetry. Changes in 4-64 labeling patterns such as the presence of diffuse labeling, blebs or vesicles, invaginations, or membrane shrinkage can be informative in characterizing mutants that impact the distribution or biosynthesis of lipids.
Beyond staining cells, determining the function of proteins participating in essential processes is necessary. The characterization of essential proteins is technically challenging because it is not possible to delete essential genes and study the phenotypic consequences. Thus, alternative approaches that deplete the protein have emerged. For example, an essential gene can be put under the control of an inducible promoter rather than its native promoter. Inducible promoters are responsive to small molecules such as; zinc16, isopropyl &#946;-d-1-thiogalactopyranoside (IPTG)17,18,19,20,21, arabinose22, vanillate17,23, and xylose23, thus the transcription of the target gene ceases and the protein of interest is depleted when the inducer is removed. Alternative approaches for depleting essential proteins of interest include synthetic riboswitches24 which use small molecule-RNA interactions to hinder transcription of target genes, CRISPR interference25,26 to block transcription of target genes, and inducible protein degradation27,28 which uses peptide tags to target proteins for degradation by the ClpXP protease. Depletion strains provide only a short time for characterization before the cells lose viability, therefore, microscopic imaging of cells over time during protein depletion is a powerful approach for characterization. Indeed, microscopy of living bacterial cells has enabled researchers to gain insights into fundamental biological processes, including the mechanisms of cell shape maintenance, secretion, and compartmentalization29.
A. tumefaciens is a bacterial plant pathogen30 and natural genetic engineer31,32. Thus, mechanisms related to pathogenicity, including host-pathogen interactions33,34,35, secretion36, and host transformation30,31,37 have been extensively investigated. To design strategies that prevent A. tumefaciens mediated disease or enhance plant transformation, the processes essential for A. tumefaciens survival need to be better understood. The use of target-specific dyes and the recent development of a protein depletion strategy for A. tumefaciens18 provides a means to investigate essential processes.
Here, detailed protocols for microscopic analysis of wildtype, mutant, and protein depletion strains of A. tumefaciens are provided. The first two protocols describe how to prepare cells and label them with target-specific dyes. The third protocol provides step-by-step directions for preparing agarose pads (Figure 1) and imaging the bacterial cells (Figure 2, Figure 3, Figure 4). These protocols may also be suitable for other bacteria with additional adaptations to account for different media conditions, growth rates, oxygen requirements, and cell structures.

<table>
	<tbody>
		<tr>
			<td>Bacterial Strains</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Agrobacterium tumefaciens C58</td>
			<td>ATCC</td>
			<td>33970</td>
			<td>Watson B, Currier TC, Gordon MP, Chilton MD, Nester EW. 1975. Plasmid required for virulence of Agrobacterium tumefaciens. J Bacteriol 123:255-264.</td>
		</tr>
		<tr>
			<td>Agrobacterium tumefaciens C58&#916;tetRA::mini-Tn7T-GM-Ptac-ctrA &#916;ctrA</td>
			<td></td>
			<td></td>
			<td>Figueroa-Cuilan W, Daniel JJ, Howell M, Sulaiman A, Brown PJB. 2016. Mini-Tn7 insertion in an artificial attTn7 site enables depeltion of the essentail master regulator CtrA in the phytopathogen Agrobacterium tumefaciens. Appl Environ Microbiol. 82:5015-5025.</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Media Components</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ATGN Minimal Medium</td>
			<td></td>
			<td></td>
			<td>To 1 L of sterilized water add 50 ml 20X Buffer, 50 ml 20X Salts, 12.5 ml 40% glucose. For plates, add 15 g Bacto Agar to 1 L of water and autoclave. Cool to 55 &#176;C and add 50 ml 20X Buffer, 50 ml 20X Salts, 12.5 ml 40% glucose.</td>
		</tr>
		<tr>
			<td>20X AT Buffer</td>
			<td></td>
			<td></td>
			<td>Add 214 g/L KH2PO4 to water and adjust pH to 7.0 with sodium hydroxide. Autoclave.</td>
		</tr>
		<tr>
			<td>NaOH</td>
			<td>Fisher BioReagents</td>
			<td>BP359</td>
			<td></td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>Fisher Chemical</td>
			<td>P288</td>
			<td></td>
		</tr>
		<tr>
			<td>20X AT Salts</td>
			<td></td>
			<td></td>
			<td>Add 40 g/L (NH4)2SO4, 3.2 g/L MgSO4&#8226;7H2O, 0.2 g/L CaCl2&#8226;2H2O, and 0.024 g/L MnSO4&#8226;H2O to water. Autoclave.</td>
		</tr>
		<tr>
			<td>(NH4)2SO4</td>
			<td>Fisher Chemical</td>
			<td>A701</td>
			<td></td>
		</tr>
		<tr>
			<td>MgSO4&#8226;7H2O</td>
			<td>Fisher BioReagents</td>
			<td>BP213</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2&#8226;2H2O</td>
			<td>Fisher BioReagents</td>
			<td>BP510</td>
			<td></td>
		</tr>
		<tr>
			<td>MnSO4&#8226;H2O</td>
			<td>Fisher Chemical</td>
			<td>M114</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Fisher Chemical</td>
			<td>D16</td>
			<td>Prepare 40% stock in water. Filter sterilize.</td>
		</tr>
		<tr>
			<td>Bacto Agar</td>
			<td>Fisher BioReagents</td>
			<td>BP1423</td>
			<td>Add 15 g to 1 L of water when preparing plates.</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Optional Media Additives</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Kanamycin</td>
			<td>GoldBio</td>
			<td>K-120</td>
			<td>Prepare as a 100 mg/ml stock solution in water and filter sterilize. Use at final concentration of 200 &#181;g/ml.</td>
		</tr>
		<tr>
			<td>IPTG</td>
			<td>GoldBio</td>
			<td>I2481C5</td>
			<td>Prepare as a 1 M stock solution in water and filter sterilize. Use at final concentration of 1 mM as needed for induction.</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Microscopy Materials</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Slides</td>
			<td>Fisherbrand</td>
			<td>12-550D</td>
			<td>25 X 75 X 1.0 mm. Clean with Sparkle glass cleaner.</td>
		</tr>
		<tr>
			<td>Microscope Cover Glass</td>
			<td>Fisherbrand</td>
			<td>12-541-B</td>
			<td>22 X 22 mm. No. 1.5. Clean with Sparkle glass cleaner.</td>
		</tr>
		<tr>
			<td>Sparkle Glass Cleaner</td>
			<td>Home Depot</td>
			<td>203261385</td>
			<td>Ammonia and alcohol free.</td>
		</tr>
		<tr>
			<td>Ultra Pure Agarose</td>
			<td>Invitrogen</td>
			<td>16500-100</td>
			<td>Add to water, PBS, or media to a final concentration of 1 - 1.5%. Melt in microwave and place on 70 C</td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Fisher BioReagents</td>
			<td>BP399500</td>
			<td>10X solution to be diluted to 1X with sterile water.</td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>Bemis</td>
			<td>PM-999</td>
			<td>Laboratory film used as gasket in agarose pad preparation.</td>
		</tr>
		<tr>
			<td>VALAP</td>
			<td></td>
			<td></td>
			<td>Add equal weights of lanolin, parafin wax, and petroleum jelly to a conical tube. Heat tube in 70 &#176;C dry, bead or water bath to melt and mix. Apply VALAP while still molten.</td>
		</tr>
		<tr>
			<td>Lanolin Butter</td>
			<td>SAAQIN</td>
			<td>SQ-LAB-R1</td>
			<td></td>
		</tr>
		<tr>
			<td>Petroleum Jelly</td>
			<td>Target Corp.</td>
			<td>06-17644</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraffin Wax</td>
			<td>Crafty Candles</td>
			<td>263012</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Target-specific dyes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Fisher BioReagents</td>
			<td>BP231-1</td>
			<td>Use to dilute stock solutions of dyes as needed.</td>
		</tr>
		<tr>
			<td>FDAAs (NADA, HADA,TADA)</td>
			<td></td>
			<td></td>
			<td>FDAAs can be synthesized or acquired through agreement with Mike VanNieuwenhze (Indiana University). Prepare 100 mM stock solution in DMSO. Use at a final concentration of 5 mM.</td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>ThermoFisher Scientific</td>
			<td>62247</td>
			<td>Prepare 1 mg/ml stock solution in DMSO. Use at final concentration of 1 &#181;g/ml.</td>
		</tr>
		<tr>
			<td>SYTOX Orange Nucleic Acid Stain</td>
			<td>Invitrogen</td>
			<td>S11368</td>
			<td>Stock concentration is 5 mM in DMSO. Use at final concentration of 5 &#181;M.</td>
		</tr>
		<tr>
			<td>FM4-64</td>
			<td>Invitrogen</td>
			<td>T3166</td>
			<td>Prepare 8 mg/ml stock solution in DMSO. Use at final concentration of 8 &#181;g/ml.</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dry bath</td>
			<td>Sheldon Manufacturing, Inc.</td>
			<td>52120-200</td>
			<td></td>
		</tr>
		<tr>
			<td>Metallic thermal beads</td>
			<td>Lab Armor</td>
			<td>42370-002</td>
			<td></td>
		</tr>
		<tr>
			<td>Epifluorescence microscope equipped with an EMCDD camera</td>
			<td></td>
			<td></td>
			<td>Nikon Eclipse TiE equipped with a QImaging Rolera em-c2 1K electron-multiplying charge-coupled-device (EMCCD) camera is used in this work.</td>
		</tr>
	</tbody>
</table>

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.

Target-specific labeling of wildtype A. tumefaciens cells 
In order to illustrate that cell morphology is not impacted by the wash steps or treatment with 1% DMSO (which is used to dilute the fluorescent dyes), cells were imaged directly from culture (Figure 2A, far left panel), after washing the cells by centrifugation as described in 1.2 (Figure 2A, left panel), o...

Watch this Scientific Journal Video about Live Cell Fluorescence Microscopy to Observe Essential Processes During Microbial Cell Growth at JoVE.com

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

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 ...

live-cell-fluorescence-microscopy-to-observe-essential-processes

Research

JoVE Journal

Biology

15.6K Views.  University of Missouri. 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.

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

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique advantage of FSM is its ability to capture the movement and turnover kinetics (assembly/disassembly) of the F-actin network within living cells. This technique is particularly useful in deriving quantitative measurements of F-actin dynamics when paired with computer vision software (qFSM).  We describe  the selection, microinjection and visualization of fluorescent actin probes in living cells. Importantly, similar procedures are applicable to visualizing other macomolecular assemblies. FSM has been demonstrated for microtubules, intermediate filaments, and adhesion complexes.  

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy FSM

Selection, microinjection, and imaging of fluorescently-labeled F-actin via fluorescent speckle microscopy (FSM).

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique ...

Time-lapse Imaging of Mitosis After siRNA Transfection

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

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

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

Cell Electrofusion Visualized with Fluorescence Microscopy

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

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

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

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

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 ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Visualizing Stromule Frequency with Fluorescence Microscopy

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

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

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

Examination of Mitotic and Meiotic Fission Yeast Nuclear Dynamics by Fluorescence Live-cell Microscopy

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does not interfere with cell physiology, and requires minimal experimental handling. The low levels of technical interference enable researchers to study cells across multiple cycles of mitosis and to observe meiosis from beginning to end. Using fluorescent tags such as Green Fluorescent Protein (GFP) and Red Fluorescent Protein (RFP), researchers can analyze different factors whose functions are important for processes like transcription, DNA replication, cohesion, and segregation. Coupled with data analysis using Fiji (a free, optimized ImageJ version), live-cell imaging offers various ways of assessing protein movement, localization, stability, and timing, as well as nuclear dynamics and chromosome segregation. However, as is the case with other microscopy methods, live-cell imaging is limited by the intrinsic properties of light, which put a limit to the resolution power at high magnifications, and is also sensitive to photobleaching or phototoxicity at high wavelength frequencies. However, with some care, investigators can bypass these physical limitations by carefully choosing the right conditions, strains, and fluorescent markers to allow for the appropriate visualization of mitotic and meiotic events.

Here, we present live-cell imaging which is a non-toxic microscopy method that allows researchers to study protein behavior and nuclear dynamics in living fission yeast cells during mitosis and meiosis.

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does ...

Live-Cell Imaging of the Life Cycle of Bacterial Predator Bdellovibrio bacteriovorus using Time-Lapse Fluorescence Microscopy

Bdellovibrio bacteriovorus is a small gram-negative, obligate predatory bacterium that kills other gram-negative bacteria, including harmful pathogens. Therefore, it is considered a living antibiotic. To apply B. bacteriovorus as a living antibiotic, it is first necessary to understand the major stages of its complex life cycle, particularly its proliferation inside prey. So far, it has been challenging to monitor successive stages of the predatory life cycle in real-time. Presented here is a comprehensive protocol for real-time imaging of the complete life cycle of B. bacteriovorus, especially during its growth inside the host. For this purpose, a system consisting of an agarose pad is used in combination with cell-imaging dishes, in which the predatory cells can move freely beneath the agarose pad while immobilized prey cells are able to form bdelloplasts. The application of a strain producing a fluorescently tagged &#946;-subunit of DNA polymerase III further allows chromosome replication to be monitored during the reproduction phase of the B. bacteriovorus life cycle.

Live-Cell Imaging of the Life Cycle of Bacterial Predator Bdellovibrio bacteriovorus using Time-Lapse Fluorescence Microscopy

Presented here is a protocol that describes monitoring of the complete life cycle of predatory bacterium Bdellovibrio bacteriovorus using time-lapse fluorescence microscopy in combination with an agarose pad and cell-imaging dishes.

Bdellovibrio bacteriovorus is a small gram-negative, obligate predatory bacterium that kills other gram-negative bacteria, including harmful pathogens. ...

Quantitative Analysis of Aspergillus nidulans Growth Rate using Live Microscopy and Open-Source Software

It is well established that colony growth of filamentous fungi, mostly dependent on changes in hyphae/mycelia apical growth rate, is macroscopically estimated on solidified media by comparing colony size. However, to quantitatively measure the growth rate of genetically different fungal strains or strains under different environmental/growth conditions (pH, temperature, carbon and nitrogen sources, antibiotics, etc.) is challenging. Thus, the pursuit of complementary approaches to quantify growth kinetics becomes mandatory in order to better understand fungal cell growth. Furthermore, it is well-known that filamentous fungi, including Aspergillus spp., have distinct modes of growth and differentiation under sub-aerial conditions on solid media or submerged cultures. Here, we detail a quantitative microscopic method for analyzing growth kinetics of&#160;the model fungus Aspergillus nidulans, using live imaging in both submerged cultures and solid media. We capture images, analyze, and quantify growth rates of different fungal strains in a reproducible and reliable manner using an open source, free software for bio-images (e.g., Fiji), in a way that does not require any prior image analysis expertise from the user.

Quantitative Analysis of Aspergillus nidulans Growth Rate using Live Microscopy and Open-Source Software

We present a label-free live imaging protocol using transmitted light microscopy techniques to capture images, analyze and quantify growth kinetics of the filamentous fungus A. nidulans in both submerged cultures and solid media. This protocol can be used in conjunction with fluorescence microscopy.

It is well established that colony growth of filamentous fungi, mostly dependent on changes in hyphae/mycelia apical growth rate, is macroscopically ...