This work was supported by NIH Grants 068625, NSF Grants 1063188 and Center of the Physics of Living Cells 0822613. Special thanks go to Dr. Marina Marjanovic in Beckman Institute for Advanced Science and Technology for the gift of rabbit eyes.

Ethics Statement: The cornea tissue from rabbits was collected in accordance with the University of Illinois Institutional Animal Care and Use guidelines.
1. TIRFM Setup
NOTE: Wear laser-safety goggles all the time.
<ol>
	<li>Make sure that all necessary optical components listed in the List of Materials are available and ready for alignment. If needed, use substitutes with equivalent functions from other companies. Ensure that Mirrors and lenses should have anti-reflective (AR) coatings matching the laser in use.</li>
	<li>Set the heights of all the optical components to the height of the center of the microscope back port.</li>
	<li>Mount the laser, laser shutter and ND filter(s). Use ND filters to attenuate the laser power down as low as possible while keeping the beam visible. Tighten screws with appropriate hex keys.</li>
	<li>Plan a beam path and mark it with tape or marker on the optical table (dotted blue lines in Figure&#160;1). For simplicity, keep straight paths along lines of holes on the optical table.</li>
	<li>Place mirror M1 (Figure&#160;1) at the first right angle turn. Place the two irises along the second straight section of the planned beam path. Adjust both the position and the tilt of M1 such that the laser goes through the irises.</li>
	<li>Place mirror M2 (Figure&#160;1) at the second right angle turn. Place the 10X beam expander (L1 &#38; L2, Figure&#160;1) along the third straight section of the path. Adjust its tilt such that the beam expander is parallel to both the optical table and the planned beam path.
	<ol>
		<li>Adjust M1 and M2 iteratively such that the laser goes straightly through the centers of both lenses of the beam expander. Repeat this step until the beam profile of the expanded beam is non-clipped Gaussian.</li>
	</ol>
	</li>
	<li>Adjust M1 to center the beam on L1 (Figure&#160;1) and M2 to center the beam on L2 (Figure&#160;1) iteratively. A bad beam profile usually means the laser is clipped; use a piece of white paper to block the beam after the beam expander to check the beam profile with eyes. For high-precision analysis, use an optical beam profiler.</li>
	<li>Adjust the distance between L1 and L2 such that the beam is collimated. Repeat Step 1.8 and 1.9 if necessary. 
	NOTE: When the beam size does not change with distance, the beam is collimated enough for a TIRFM setup. To further improve the beam collimation, tools such as a shearing interferometer could be used. A typical beam size after expansion is ~20 mm.</li>
	<li>Shutter the laser. Unscrew the microscope objective and screw in a fluorescent alignment target. Place mirrors M3 and M4 (Figure&#160;1) to direct the expanded beam into the microscope port and onto the dichroic mirror inside the turret. Ensure that the laser beam bounces off the dichroic and towards the ceiling.</li>
	<li>Adjust M3 to center the brightest part of the beam on the fluorescent target, and M4 to adjust the beam tilt to be vertical.</li>
	<li>Shutter the laser and screw the objective back in. If the alignment in the previous step is done well there should be a symmetric spot exiting the objective. Fine-tune the tilts of M3 and M4 to optimize the laser power and beam profile out of the objective.</li>
	<li>Mount an EMCCD camera to the microscope and connect the camera to a computer. Start the software for the camera.</li>
	<li>Mount a fluorescent sample (solution of fluorophores) on the microscope. Look at the bright fluorescent spot on the camera. Check that the spot does not shift on the screen as the focus is changed.</li>
	<li>Place the TIR lens (L3, Figure&#160;1) on the XYZ translation stage at a distance from the back focal plane of the objective which is equal to the focal length of L3 (~30 cm). Adjust the position of L3 such that the laser goes through the center of the lens.</li>
	<li>Translate L3 along the beam path to adjust the beam collimation. Make sure that the beam is still centered on the monitor and symmetrical in shape. 
	NOTE: The illumination area at the sample plane increases with decreasing TIR lens focal length. In general, use the smallest focal length that could fit on the set-up.</li>
	<li>Translate L3 perpendicular to the beam path to tilt the beam out of the objective. Keep translating the TIR lens such that TIR is achieved. Observe a sample of fluorescent beads through the EMCCD camera, and fine-tune L3 to get good SNR.</li>
</ol>
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51774/51774fig1highres.jpg" width="500" /> 
Figure 1. Optical configuration for total internal reflection fluorescence microscopy (TIRFM).
2. FIONA on Cy3-DNA
<ol>
	<li>Clean microscope slides and coverslips: rinse microscope slides and coverslips with ddH2O and isopropanol and dry them with nitrogen gas; place the slides and coverslips in the plasma cleaner for 5 min under argon plasma.</li>
	<li>Construct sample chambers (as sketched in Figure&#160;2).
	<ol>
		<li>Place a tissue on the bench and then put a piece of lens paper on top of the tissue. Place the slide on the lens paper. Make sure that the clean side of the slide is upwards.</li>
		<li>Apply two strips of double-sided tape onto the slide along the long edges, leaving a gap of 3-5 mm at the center. Place the cleaned coverslip on top of the slide. Make sure that the clean side of the coverslip is facing the slide.</li>
		<li>Use a pipette tip to press down over the double-sided tape. Use a razor to remove excess tape from the slide so that the tape remains only under the coverslip. 
		NOTE: The open ends of the chamber are left open and serve as inlet and outlet. The chamber volume is several microliters.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51774/51774fig2highres.jpg" width="500" /> 
Figure 2. Sketch of a typical sample chamber. (a) Top view; (b) Side view from the right; (c) Side view from the front.
<ol start="3">
	<li>Immobilize Cy3-DNA on inner surfaces of the sample chambers.
	<ol>
		<li>Prepare T-50 buffer (10 mM Tris-HCl pH 8.0, 50 mM NaCl). Prepare BSA-biotin in T-50 at a final concentration of 1 mg/ml. Prepare T50-BSA buffer by dissolving BSA in T-50 at a final concentration of 10 mg/ml.</li>
		<li>Prepare 0.5 mg/ml neutravidin in T50-BSA buffer. Prepare biotinylated Cy3 labeled DNA (Cy3-DNA) in T50-BSA at a final concentration of 5-10 pM.</li>
		<li>Pipette 10 &#181;l BSA-biotin (1 mg/ml) into the sample chamber. Wait for 5 min.</li>
		<li>Wash the chamber with 40 &#181;l T50-BSA. Pipette 10 &#181;l neutravidin (0.5 mg/ml) into the sample chamber. Incubate for 5 min.</li>
		<li>Wash the chamber with 40 &#181;l T50-BSA. Pipette 20 &#181;l Cy3-DNA into the sample chamber. Incubate for 5 min and then wash the chamber with 80 &#181;l T50-BSA.</li>
	</ol>
	</li>
	<li>Image Cy3-DNA single molecules under TIRFM.
	<ol>
		<li>Prepare imaging buffer (100 &#181;l) by mixing 1 &#181;l Protocatechuate-3,4-dioxygenase (PCD, 5 &#181;M), 4 &#181;l Protocatechuic acid (PCA, 62.5 mM), 50 &#181;l 6-hydroxy-2,5,7,8-tetramethylchromane-2-carboxylic acid (Trolox, 2 mM in T-50), and 45 &#181;l T50-BSA.</li>
		<li>Pipette in 30 &#181;l imaging buffer and wait for 8-10 min.</li>
		<li>Mount the sample for imaging on a TIRFM that is equipped with a green laser (532 nm), a 100X oil immersion objective (1.45 NA), and an EMCCD camera.</li>
		<li>Set the exposure time to 100 to 500 msec and EM gain to 25 to 100. Acquire a movie of the sample for 1,000 frames.</li>
	</ol>
	</li>
	<li>Perform FIONA data analysis on the recorded images of Cy3-DNA.
	<ol>
		<li>Determine the effective pixel size (i.e., conversion factor from pixels to nanometers) by dividing the physical pixel size (read from the specification sheet of the EMCCD camera) by the total magnification (the magnification of the microscope objective multiplied by any extra magnifications).</li>
		<li>Determine the conversion factor from pixel intensities to photon numbers by dividing CCD sensitivity (i.e., electrons per A/D count, read from the specification sheet of the CCD camera) by the electron multiplier (EM) gain used during image acquisition.</li>
		<li>Compile and run FIONA.pro for FIONA analysis. Use this IDL program to import the acquired image, to input the effective pixel size (from Step 2.5.1) and the conversion factor from intensity to photon number (from Step 2.5.2), and to choose spots for FIONA analysis. 
		NOTE: At the end, the program will output the fitting results with 2D Gaussian functions, as well as total photon numbers and localization precision. A typical result is shown in the section of representative results and Figures&#160;4c-4d.</li>
		<li>Compile and run phcount.pro to characterize photon count. Use this IDL program to measure the average number of photons emitted by a fluorophore before photobleaching, to import the acquired image and to input the conversion factor from intensity to photon number (from Step 2.5.2). 
		NOTE: The program will then detect fluorescent spots automatically (manual selection is an option), calculate photon counts as a function of frame number, and output the traces of photon counts.
		<ol>
			<li>Discard bad traces and specify frame ranges after photobleaching for baseline correction. At the end, the program will output a list of total photon numbers for all the spots not discarded. Then plot the distribution of the photon numbers and fit the distribution with an exponential decay to obtain the average photon number. A typical result is shown in the section of representative results and Figures&#160;4e-4f.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
3. FIONA Applied to Quantify Motor (e.g., Myosin on Actin) Dynamics at Nanometer Scale
<ol>
	<li>Polymerize actin (i.e. prepare F-actin) one day before FIONA experiment.
	<ol>
		<li>Reconstitute G-actin (monomer) and biotin G-actin (monomer) to 10 mg/ml with general actin buffer. Stir to make sure both are fully dissolved, and keep both on ice.</li>
		<li>Mix 10 &#181;l G-actin (monomer) with 1.7 &#181;l biotin G-actin in a 1.5 ml microcentrifuge tube. Add 100 &#181;l ice cold actin polymerization buffer.</li>
		<li>Leave the mixture overnight at 4 &#176;C (F-actin formed) and then add ddH2O to a total volume of 1 ml.</li>
		<li>Store the actin filaments (F-actin) at 4 &#176;C for later use in experiments. 
		NOTE: Filaments will disintegrate and shorten over time, but can be used for at least two weeks.</li>
	</ol>
	</li>
	<li>Prepare sample for imaging.
	<ol>
		<li>Make a sample chamber (as described in Protocol 2.1 and 2.2). Pipette in 20 &#181;l biotinylated BSA at 1 mg/ml in ddH2O. Incubate for 10 min. Rinse with 30 &#181;l ddH2O. 
		NOTE: This blocks the glass surface and lays down biotin for binding of the actin filaments. Biotinylated poly-L-lysine &#8211; polyethylene glycol (PLL-PEG) could serve the same function.</li>
		<li>Pipette in 0.5 mg/ml neutravidin. Incubate for 2 min and then wash the chamber with 30 &#181;l M5 buffer.</li>
		<li>Pipette in the prepared F-actin diluted 25 times in general actin buffer, to final concentration ~0.004 mg/ml. Wait 10 min, rinse the chamber with 30 &#181;l buffer.</li>
		<li>Dilute myosin Va with FLAG tags 30-fold in M5 buffer (20 mM HEPES (pH 7.6), 2 mM MgCl2, 25 mM KCl, 1 mM EGTA) to a final concentration of 250 nM. Mix 1 &#181;l myosin with 1 &#181;l Anti-FLAG-Qdot705 (~1 &#181;M, conjugated from Anti-FLAG antibodies and Qdot705 using the Qdot705 Antibody Conjugation Kit according to the instruction manual from the manufacturer). Add in 8 &#956;l M5 to fill to 10 &#181;l. Pipette up and down to mix well. Incubate for 10 min on ice. 
		NOTE: This yields a mixture of 1 motor to 4 quantum dots, at ~25 nM myosin concentration.</li>
	</ol>
	</li>
	<li>Imaging of the myosin walking on actin.
	<ol>
		<li>Prepare imaging buffer (100 &#181;l) by mixing 84 &#181;l M5-BSA (M5 buffer with 1 mg/ml BSA), 1 &#181;l ATP (50 &#181;M in ddH2O), 2 &#181;l DTT (500 mM in ddH2O), 1 &#181;l CK (500 U/ml), 5 &#181;l CP (200 mM), 1 &#181;l PCD, 4 &#181;l PCA, 1 &#181;l myosin-qdot after diluted another 10-fold to 2.5 nM myosin concentration, and 1 &#181;l BME.</li>
		<li>Pipette in 20 &#181;l imaging buffer to sample chamber and incubate for 8-10 min.</li>
		<li>Image the sample on TIRF microscope at 30 msec exposure. Acquire at least 1,000 frames. Adjust the volume of myosin-qdot in Step 3.3.1 if necessary.</li>
	</ol>
	</li>
	<li>Perform data analysis and find the step-size of myosin walking.
	<ol>
		<li>Open the video file in ImageJ17 and crop the video around a moving spot. Crop a large enough area that the spot never gets within 20 pixels of the edge, and make sure there are no other spots in the video. Ensure that this spot is moving in a linear path.</li>
		<li>Track the spot through the video to generate x and y coordinates through time, in pixels, by applying FIONA analysis (Step 2.5.3) to each and every frame of the video.</li>
		<li>Convert pixels to nanometers, as described in the previous section.</li>
		<li>Calculate displacement from initial position as a function of time.</li>
		<li>Run t-test on the displacement to obtain the steps of myosin walking. 
		NOTE: The program provided for t-test (step_t_test.zip) is coded in IDL and consists of 14 subroutines in the folder.
		<ol>
			<li>Open all of the subroutines in IDL and compile all twice. Then run mtltyanalysis_ttest.pro and choose a text file containing only the distance data in a single column. An output Excel file will be generated, containing the raw data, the fit, and the step size.</li>
		</ol>
		</li>
		<li>Delete all zero-values from the step-size column. Plot the distribution of the step sizes using Origin or MATLAB. Fit a Gaussian to the histogram. 
		NOTE: Step sizes of zero-values need to be deleted because a step of zero means that no step is taken from the previous frame. The fitting gives a peak around 36 nm (as shown in Figure&#160;5).</li>
	</ol>
	</li>
</ol>
4. Thick Sample Preparation for FIONA
<ol>
	<li>Prepare quantum dots encapsulated in sol-gel.
	<ol>
		<li>Mix 4.5 ml TMOS, 1 ml ddH2O and 100 &#181;l HCl (120 mM). Sonicate the mixture on ice for 30 min in the ultrasonic cleaner specified in the List of Materials (frequency = 40 kHz, heater = off). Mix the solution every 10 min.</li>
		<li>Dilute 1.5 &#181;l Qdot605 in 1.5 ml HEPES (50 mM pH 7.2). Mix solution with 1.5 ml TMOS from the previous step.</li>
		<li>Pour the mixture into a glass bottom dish. Seal the glass bottom dish with Parafilm and store at 4 &#186;C for 1. 5 hr.</li>
		<li>Add 2 ml 1% BME in PBS to the sample dish and incubate at room temperature for 30 min before imaging.</li>
	</ol>
	</li>
	<li>Prepare cornea sample stained with quantum dots. 
	NOTE: Rabbit-eyes were gifts from Dr. Marina Marjanovic.
	<ol>
		<li>Separate the cornea from the eyes and cut it into 3 mm x 3 mm pieces.</li>
		<li>Dilute 1 &#181;l Qdot 605-streptavidin in 1 ml of PBS. Incubate the cornea tissue with 1 nM Qdots solution at 4 &#186;C for 1 hr. Wash the tissue with PBS.</li>
		<li>Take a clean glass slide and a #1.5 coverslip. Put 4 layers of double sided tape on the glass slide along the longer side, and put another 4 layers of double sided tape parallel to the previous tape and leave a channel about 1 cm in between. Put the tissue from the previous step in the middle of the channel, and cover it with a coverslip.</li>
		<li>Press gently on the sides of the coverslip to make it stick to the tapes. Wet the channel with 50 &#181;l PBS before imaging.</li>
	</ol>
	</li>
	<li>Image quantum dots in sol-gel and cornea.
	<ol>
		<li>For FIONA imaging in thick samples, use a 60X water-immersion objective with working distance of 0.27 mm, or a 60X water-immersion objective with working distance of 0.28 mm.</li>
		<li>Mount the sample on the microscope. Adjust the TIRF lens such that it reaches the epi-fluorescence mode (i.e. the laser beam comes out of the objective with an angle against the coverslip). Insert an extra magnification lens (3.3X or 4.0X).</li>
		<li>Move the focal plane of the objective to a desired z position (e.g. &#62;200 &#181;m). Record images of immobile quantum dots in the sample.</li>
	</ol>
	</li>
	<li>Perform FIONA analysis as described in section 2.5 of this protocol.</li>
</ol>

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The authors declare that they have no competing financial interests.

FIONA is a technique to localize the position of a fluorescent emitter (organic fluorophore or quantum dot) with nanometer precision and temporal resolution down to 1 msec4-8. When enough photons are collected, this technique allows to determine the position of a fluorescent emitter much more accurately than the diffraction limit (~200 nm) and thus this technique opens a way to observe what has not been seen with conventional/traditional optical microscopy4-8. Since its inventi...

Around 1882, Ernst Abbe found that the resolution of a visible light microscope is ~&#955;/2NA, or ~200 nm (where &#955; is the wavelength and NA is the numerical aperture)1,2. Therefore any object smaller than this dimension would appear as a diffraction-limited spot in an optical microscope. However, it is possible to determine the center of the spot, that is, the location of the object, with a much higher precision3. Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in the x-y plane4. The precision of localization,&#160;&#963;&#181; (i.e., the standard error of the mean), depends on the total number of collected photons, <img alt="Equation 1" fo:content-width="1in" src="/files/ftp_upload/51774/intro_eqn.jpg" width="100" />, where N is the photon count, s is the standard deviation of the fluorescent spot, a is the pixel size of the imaging detector, and b is the standard deviation of the background3,4. For a fluorophore emitting ~ 10,000 photons, FIONA can achieve ~1 nm precision4.
FIONA can be used to accurately determine the position of a stationary emitter, or a moving one (assuming images can be taken fast enough). FIONA can be applied sequentially to the frames of the movie and thus track the motion of the single molecule4-8. Photo-protective reagents may be necessary to ensure that the sample does not photodegrade. Furthermore, the fluorescent object itself may be of any size, smaller or larger than the diffraction limit&#8212;e.g., it may consist of an organelle (~1 &#181;m) with many fluorescent proteins dispersed on its membrane. Using FIONA can still yield a very accurate (nanometer) average of its average center-of-mass. The great improvement in localization precision by FIONA allows resolving nanometer-scale movements over time. This has pushed microscopy into the molecular length scale4-8.
Since its invention, variants of FIONA have been developed. For example, bright-field imaging with one-nanometer accuracy (bFIONA)9, a slight variant of FIONA, images and localizes dense objects such as melanosomes in vivo (dark objects containing the pigment melanin) with transmitted light. In addition, FIONA has been employed to resolve multiple dyes. For example, single-molecule high-resolution imaging with photobleaching (SHRImP)10,11 or single-molecule high-resolution colocalization (SHREC)12 have been developed to resolve two dyes within about 10 nm. (Notice that this is resolution, i.e. how accurately one can tell identical dyes apart.) More recently, FIONA analysis has contributed to the localization process of certain super-resolution microscopy such as stochastic optical reconstruction microscopy (STORM)13-15 and photo-activated localization microscopy (PALM)16, in which temporary dark fluorophores are excited, and then the fluorescence is localized. By repeatedly exciting a fairly low density of dyes (less than one per diffraction limited spot), and then collecting the fluorescence, analyzing each of them by FIONA, one can build up a high-resolution map. The resolution is then just limited by the number of photons each dye puts out, as well as things like keeping the sample stationary (including, e.g., the microscope stage) during the acquisition.
In this paper, a summary of the FIONA technique and briefly describe examples of research that have been performed using FIONA is reported. First, how to set up the required equipment for FIONA experiments, i.e., a total internal reflection fluorescence microscopy (TIRFM), with details on aligning the optics, is described. Then how to carry out a simple FIONA experiment on localizing immobilized Cy3-DNA single molecules using appropriate protocols, is illustrated. After that, the use of FIONA to measure the 36 nm step size of a single truncated myosin Va motor labeled with a quantum dot is presented. Myosin Va is an essential processive motor protein which carries cellular cargo while translocating along actin filaments. Here a myosin Va construct truncated is used to remove domains irrelevant to the step size, and with a FLAG tag added to the C-terminus to allow ease of labeling with quantum dots functionalized with Anti-FLAG antibodies. This experiment is done under low ATP to slow down the myosin and allow the use of long enough exposure times to get a good photon count in every frame. Any sufficiently bright fluorescent label could be substituted in the following protocol. Lastly, recent effort of extending the application of FIONA to thick samples is reported. As a proof-of-principle, quantum dots were soaked in sol-gels and rabbit eye corneas and then imaged and localized using FIONA. For imaging, a 60X water immersion objective with NA=1.2 was used because this objective has a longer working distance than previously used 100X oil immersion objective. To compensate the loss in the magnification in the objective, an extra-magnification lens (3.3X or 4.0X) was inserted in the emission path. In addition, epi-fluorescence (not TIR) microscopy needs to be used to access deep regions in the thick samples. It is shown that quantum dots soaked deep in sol-gels and in rabbit eye corneas (Z &#62; 200 &#181;m) can be localized with 2-3 nm precision.

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			<td height="21" style="height:21px;width:220px;">NaCl</td>
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			<td>Protocatechuic acid&#160;</td>
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			<td>Tetramethyl orthosilicate</td>
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			<td>Quantity: 2</td>
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			<td height="21" style="height:21px;width:220px;">XYZ translation stage, &#189;&#8221; travel&#160;</td>
			<td style="width:211px;">Thorlabs&#160;</td>
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			<td height="21" style="height:21px;width:220px;">Laser</td>
			<td style="width:211px;">World Star Technologies&#160;</td>
			<td style="width:211px;">TECGL-30</td>
			<td>532 nm, 30 mW</td>
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fluorescence imaging with one-nanometer accuracy (fiona)

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in the x-y plane. Here a summary of the FIONA technique is reported and examples of research that have been performed using FIONA are briefly described. First, how to set up the required equipment for FIONA experiments, i.e., a total internal reflection fluorescence microscopy (TIRFM), with details on aligning the optics, is described. Then how to carry out a simple FIONA experiment on localizing immobilized Cy3-DNA single molecules using appropriate protocols, followed by the use of FIONA to measure the 36 nm step size of a single truncated myosin Va motor labeled with a quantum dot, is illustrated. Lastly, recent effort to extend the application of FIONA to thick samples is reported. It is shown that, using a water immersion objective and quantum dots soaked deep in sol-gels and rabbit eye corneas (&#62;200 &#181;m), localization precision of 2-3 nm can be achieved.

A typical objective-type TIRFM setup is shown in Figure&#160;3. First, surface-immobilized Cy3-DNA sample was imaged. A typical image is shown in Figure&#160;4a. The image was taken with exposure time 0.5 sec, with EM gain = 50 and CCD sensitivity = 12.13 for the camera. The point-spread-function (PSF) of a single Cy3-DNA molecule is shown in Figure&#160;4b (from the spot indicated by the arrow in Figure&#160;4a), where the color-bar shows the scale of p...

Watch this Scientific Journal Video about Fluorescence Imaging with One-nanometer Accuracy FIONA at JoVE.com

Fluorescence Imaging with One-nanometer Accuracy FIONA

Single fluorophores can be localized with nanometer precision using FIONA. Here a summary of the FIONA technique is reported, and how to carry out FIONA experiments is described.

Fluorescence Imaging with One-nanometer Accuracy (FIONA)

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in ...

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17.5K Views.  University of Illinois at Urbana-Champaign. Single fluorophores can be localized with nanometer precision using FIONA. Here a summary of the FIONA technique is reported, and how to carry out FIONA experiments is described.

Video: Fluorescence Imaging with One-nanometer Accuracy FIONA

Born Normalization for Fluorescence Optical Projection Tomography for Whole Heart Imaging

Optical projection tomography is a three-dimensional imaging technique that has been recently introduced as an imaging tool primarily in developmental biology and gene expression studies. The technique renders biological sample optically transparent by first dehydrating them and then placing in a mixture of benzyl alcohol and benzyl benzoate in a 2:1 ratio (BABB or Murray s Clear solution). The technique renders biological samples optically transparent by first dehydrating them in graded ethanol solutions then placing them in a mixture of benzyl alcohol and benzyl benzoate in a 2:1 ratio (BABB or Murray s Clear solution) to clear. After the clearing process the scattering contribution in the sample can be greatly reduced and made almost negligible while the absorption contribution cannot be eliminated completely. When trying to reconstruct the fluorescence distribution within the sample under investigation, this contribution affects the reconstructions and leads, inevitably, to image artifacts and quantification errors.. While absorption could be reduced further with a permanence of weeks or months in the clearing media, this will lead to progressive loss of fluorescence and to an unrealistically long sample processing time. This is true when reconstructing both exogenous contrast agents (molecular contrast agents) as well as endogenous contrast (e.g. reconstructions of genetically expressed fluorescent proteins).

We suggest a Born normalized approach for Optical Projection Tomography (BnOPT) that accounts for the absorption properties of imaged samples to obtain accurate and quantitative fluorescence tomographic reconstructions. We use the proposed algorithm to reconstruct the fluorescence molecular probe distribution within small animal organs.

Optical projection tomography is a three-dimensional imaging technique that has been recently introduced as an imaging tool primarily in developmental ...

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Visualizing developing organ formation as well as progession and treatment of disease often heavily relies on the ability to optically interrogate molecular and functional changes in intact living organisms. Most existing optical imaging methods are inadequate for imaging at dimensions that lie between the penetration limits of modern optical microscopy (0.5-1mm) and the diffusion-imposed limits of optical macroscopy (&gt;1cm) [1]. Thus, many important model organisms, e.g. insects, animal embryos or small animal extremities, remain inaccessible for in-vivo optical imaging. 
Although there is increasing interest towards the development of nanometer-resolution optical imaging methods, there have not been many successful efforts in improving the imaging penetration depth. The ability to perform in-vivo imaging beyond microscopy limits is in fact met with the difficulties associated with photon scattering present in tissues. Recent efforts to image entire embryos for example [2,3] require special chemical treatment of the specimen, to clear them from scattering, a procedure that makes them suitable only for post-mortem imaging. These methods however evidence the need for imaging larger specimens than the ones usually allowed by two-photon or confocal microscopy, especially in developmental biology and in drug discovery.
We have developed a new optical imaging technique named Mesoscopic Fluorescence Tomography [4], which appropriate for non-invasive in-vivo imaging at dimensions of 1mm-5mm. The method exchanges resolution for penetration depth, but offers unprecedented tomographic imaging performance and it has been developed to add time as a new dimension in developmental biology observations (and possibly other areas of biological research) by imparting the ability to image the evolution of fluorescence-tagged responses over time. As such it can accelerate studies of morphological or functional dependencies on gene mutations or external stimuli, and can importantly, capture the complete picture of development or tissue function by allowing longitudinal time-lapse visualization of the same, developing organism.
The technique utilizes a modified laboratory microscope and multi-projection illumination to collect data at 360-degree projections. It applies the Fermi simplification to Fokker-Plank solution of the photon transport equation, combined with geometrical optics principles in order to build a realistic inversion scheme suitable for mesoscopic range. This allows in-vivo whole-body visualization of non-transparent three-dimensional structures in samples up to several millimeters in size.
We have demonstrated the in-vivo performance of the technique by imaging three-dimensional structures of developing Drosophila tissues in-vivo and by following the morphogenesis of the wings in the opaque Drosophila pupae in real time over six consecutive hours.

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Mesoscopic fluorescence tomography operates beyond the penetration limits of tissue-sectioning fluorescence microscopy. The technique is based on multi-projection illumination and a photon transport description. We demonstrate in-vivo whole-body 3D visualization of the morphogenesis of GFP-expressing wing imaginal discs in Drosophila melanogaster.

Visualizing  developing organ formation as well as progession and treatment of disease often  heavily relies on the ability to optically interrogate ...

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

Time-lapse Imaging of Mitosis After siRNA Transfection

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

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

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

Cell Electrofusion Visualized with Fluorescence Microscopy

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

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

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

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

Rapid Analysis and Exploration of Fluorescence Microscopy Images

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized image analysis tools are available, most traditional image-analysis-based workflows have steep learning curves (for fine tuning of analysis parameters) and result in long turnaround times between imaging and analysis. In particular, cell segmentation, the process of identifying individual cells in an image, is a major bottleneck in this regard.
Here we present an alternate, cell-segmentation-free workflow based on PhenoRipper, an open-source software platform designed for the rapid analysis and exploration of microscopy images. The pipeline presented here is optimized for immunofluorescence microscopy images of cell cultures and requires minimal user intervention. Within half an hour, PhenoRipper can analyze data from a typical 96-well experiment and generate image profiles. Users can then visually explore their data, perform quality control on their experiment, ensure response to perturbations and check reproducibility of replicates. This facilitates a rapid feedback cycle between analysis and experiment, which is crucial during assay optimization. This protocol is useful not just as a first pass analysis for quality control, but also may be used as an end-to-end solution, especially for screening. The workflow described here scales to large data sets such as those generated by high-throughput screens, and has been shown to group experimental conditions by phenotype accurately over a wide range of biological systems. The PhenoBrowser interface provides an intuitive framework to explore the phenotypic space and relate image properties to biological annotations. Taken together, the protocol described here will lower the barriers to adopting quantitative analysis of image based screens.

Here we describe a workflow for rapidly analyzing and exploring collections of fluorescence microscopy images using PhenoRipper, a recently developed image-analysis platform.

Despite rapid advances in high-throughput microscopy, quantitative image-based assays still pose significant challenges. While a variety of specialized ...

Visualizing Stromule Frequency with Fluorescence Microscopy

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

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

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

Mixed Primary Cultures of Murine Small Intestine Intended for the Study of Gut Hormone Secretion and Live Cell Imaging of Enteroendocrine Cells

The gut is the largest endocrine organ of the body, with hormone-secreting enteroendocrine cells located along the length of the gastrointestinal epithelium. Despite their physiological importance, enteroendocrine cells represent only a small fraction of the epithelial cell population and in the past, their characterization has presented a considerable challenge resulting in a reliance on cell line models. Here, we provide a detailed protocol for the isolation and culture of mixed murine small intestinal cells. These primary cultures have been used to identify the signaling pathways underlying the stimulation and inhibition of gut peptide secretion in response to a number of nutrients and neuropeptides as well as pharmacological agents. Furthermore, in combination with the use of transgenic fluorescent reporter mice, we have demonstrated that these primary cultures become a powerful tool for the examination of fluorescently-tagged enteroendocrine cells at the intracellular level, using methods such as patch clamping and single-cell calcium and cAMP-FRET imaging.

This protocol describes the isolation and culture of mixed murine small intestinal cells. These primary intestinal cultures enable the investigation of signaling pathways underlying gut peptide secretion using a number of techniques.

The gut is the largest endocrine organ of the body, with hormone-secreting enteroendocrine cells located along the length of the gastrointestinal ...

Fluorescence Recovery after Photobleaching of Yellow Fluorescent Protein Tagged p62 in Aggresome-like Induced Structures

Fluorescence recovery after photobleaching (FRAP)&#160;is a microscopy technique that can be used to quantify protein mobility in live cells. In a typical FRAP experiment, steady-state fluorescence is observed by repeated imaging with low-intensity laser light. Subsequently, the fluorescent molecules are rapidly and irreversibly impaired via brief exposure to high-intensity laser light. Information about protein mobility is obtained by monitoring the recovery of fluorescence. We used FRAP to determine the mobility of p62 in aggresome-like induced structures (ALIS) in murine macrophages after stimulation with lipopolysaccharide (LPS). Because many existing FRAP protocols are either incomplete or complex, our goal was to provide a comprehensive, practical, and straightforward step-by-step protocol for FRAP experiments with live cells. Here, we describe RAW264.7 macrophage transfection with yellow fluorescent protein-p62 (YFP-p62), induction of ALIS by exposing the cells to LPS, and a step-by-step method for collecting prebleach and postbleach FRAP images and data analysis. Finally, we discuss important factors to consider when conducting a FRAP experiment.

We describe a comprehensive and practical protocol for fluorescence recovery after photobleaching experiments with live cells. Although the protocol was used to measure the mobility of yellow fluorescent protein-tagged p62 in aggresome-like induced structures, it can be applied to a variety of microscopy systems and fluorescent proteins.

Fluorescence recovery after photobleaching (FRAP)&#160;is a microscopy technique that can be used to quantify protein mobility in live cells. In a typical ...