Name: fluorescence imaging with one-nanometer accuracy (fiona)
Uploaded: 2014-09-26T15:00:00.000+00:00
Duration: 11 min 56 s
Description: Watch this Scientific Journal Video about Fluorescence Imaging with One-nanometer Accuracy FIONA at JoVE.com

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.

<table><tbody><tr><td>Double-sided tape</td><td>3M</td><td></td><td>~75 &amp;#181;m thick</td></tr><tr><td>EMCCD camera</td><td>Andor Technology</td><td>DU-897E-CS0-#BV</td><td></td></tr><tr><td>Ultrasonic cleaner</td><td>Branson</td><td>2510</td><td></td></tr><tr><td>Fluorescence filter set</td><td>Chroma</td><td>49016</td><td></td></tr><tr><td>Actin polymerization buffer&amp;nbsp;</td><td>Cytoskeleton&amp;nbsp;</td><td>BSA02</td><td></td></tr><tr><td>Biotin G-actin</td><td>Cytoskeleton&amp;nbsp;</td><td>AB07</td><td></td></tr><tr><td>G-actin&amp;nbsp;</td><td>Cytoskeleton&amp;nbsp;</td><td>AKL95</td><td></td></tr><tr><td>General actin buffer&amp;nbsp;</td><td>Cytoskeleton&amp;nbsp;</td><td>BSA01</td><td></td></tr><tr><td>Laser shutter (with driver)</td><td>Electro-Optical Products Corp.&amp;nbsp;</td><td>SH-10-MP</td><td></td></tr><tr><td>IDL</td><td>Exelis Visual Information Solutions</td><td></td><td></td></tr><tr><td>Neutravidin&amp;nbsp;</td><td>Fisher Scientific</td><td>PI-31000</td><td></td></tr><tr><td>Coverslip</td><td>Fisherbrand&amp;nbsp;</td><td>22X30-1.5</td><td>0.16-0.19 mm thick</td></tr><tr><td>Microscope slide</td><td>Gold Seal Microslides&amp;nbsp;</td><td>30103X1</td><td>0.93-1.05 mm thick</td></tr><tr><td>Plasma cleaner</td><td>Harrick Plasma</td><td>PDC-001</td><td></td></tr><tr><td>Glass bottom dish&amp;nbsp;</td><td>In Vitro Scientific</td><td>D35-20-1.5-N</td><td></td></tr><tr><td>Cy3-DNA oligos</td><td>Integrated DNA Technologies</td><td></td><td>5&#039;-Cy3/GCCTCGCTGCCGTCGCCA-3&#039;Bio</td></tr><tr><td>Fluorescent beads&amp;nbsp;</td><td>Invitrogen</td><td>T-7280</td><td></td></tr><tr><td>Qdot 605-streptavidin&amp;nbsp;</td><td>Invitrogen</td><td>Q10101MP</td><td></td></tr><tr><td>Qdot605&amp;nbsp;</td><td>Invitrogen</td><td>Q21301MP</td><td></td></tr><tr><td>Qdot705</td><td>Invitrogen</td><td>Q22021MP&amp;nbsp;</td><td></td></tr><tr><td>Qdot705 Antibody Conjugation Kit</td><td>Invitrogen</td><td>Q22061MP</td><td></td></tr><tr><td>MATLAB</td><td>MathWorks</td><td></td><td></td></tr><tr><td>Optical table</td><td>Newport Corp</td><td></td><td>RS4000 Series</td></tr><tr><td>60X Objective</td><td>Nikon</td><td>Plan Apo VC 60x WI</td><td></td></tr><tr><td>100X Objective</td><td>Olympus</td><td>PlanApo 100X/1.45 Oil &amp;infin;/0.17</td><td></td></tr><tr><td>60X Objective</td><td>Olympus</td><td>UPlanApo 60X/1.20W</td><td></td></tr><tr><td>Inverted microscope</td><td>Olympus</td><td>IX71/IX70/IX81</td><td></td></tr><tr><td>Origin</td><td>OriginLab</td><td></td><td></td></tr><tr><td>Anti-FLAG antibody</td><td>Sigma Aldrich</td><td>F7425-.2MG</td><td></td></tr><tr><td>ATP</td><td>Sigma Aldrich</td><td>A7699</td><td></td></tr><tr><td>BME&amp;nbsp;</td><td>Sigma Aldrich</td><td>63689-25ML-F</td><td></td></tr><tr><td>BSA</td><td>Sigma Aldrich</td><td>A7906</td><td></td></tr><tr><td>BSA-biotin</td><td>Sigma Aldrich</td><td>A8549-10MG</td><td></td></tr><tr><td>CK</td><td>Sigma Aldrich</td><td>C3755</td><td>Creatine Phosphokinase from rabbit muscle</td></tr><tr><td>CP</td><td>Sigma Aldrich</td><td>P1937</td><td>Phosphocreatine di(tris) salt</td></tr><tr><td>DTT</td><td>Sigma Aldrich</td><td>43815</td><td>DL-Dithiothreitol</td></tr><tr><td>EGTA</td><td>Sigma Aldrich</td><td>E3889</td><td>Ethylene glycol-bis(2-aminoethylether)-&lt;em&gt;N,N,N&amp;prime;,N&amp;prime;&lt;/em&gt;-tetraacetic acid</td></tr><tr><td>HCl&amp;nbsp;</td><td>Sigma Aldrich</td><td>93363-500G</td><td></td></tr><tr><td>HEPES&amp;nbsp;</td><td>Sigma Aldrich</td><td>H0887</td><td></td></tr><tr><td>KCl</td><td>Sigma Aldrich</td><td>P9333</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma Aldrich</td><td>M1028</td><td></td></tr><tr><td>NaCl</td><td>Sigma Aldrich</td><td>S7653</td><td></td></tr><tr><td>PCA</td><td>Sigma Aldrich</td><td>03930590</td><td>Protocatechuic acid&amp;nbsp;</td></tr><tr><td>PCD</td><td>Sigma Aldrich</td><td>P8279</td><td>Protocatechuate-3,4-dioxygenase&amp;nbsp;</td></tr><tr><td>TMOS</td><td>Sigma Aldrich</td><td>341436-25G</td><td>Tetramethyl orthosilicate</td></tr><tr><td>Tris-HCl&amp;nbsp;</td><td>Sigma Aldrich</td><td>93363</td><td></td></tr><tr><td>Trolox</td><td>Sigma Aldrich</td><td>238813</td><td>6-hydroxy-2,5,7,8-tetramethylchromane-2-carboxylic acid</td></tr><tr><td>1&amp;rdquo; diameter broadband dielectric mirrors with mounts</td><td>Thorlabs&amp;nbsp;</td><td>BB1-E02, KM100</td><td>Quantity: 2</td></tr><tr><td>&amp;frac12;&amp;rdquo; diameter posts</td><td>Thorlabs&amp;nbsp;</td><td>TR4</td><td>Quantity &amp;ge; 6</td></tr><tr><td>10X beam expander</td><td>Thorlabs&amp;nbsp;</td><td>BE10M-A</td><td></td></tr><tr><td>2&amp;rdquo; diameter f = 300 mm lens with mount</td><td>Thorlabs&amp;nbsp;</td><td>LA1256-A, LMR2</td><td>TIR lens</td></tr><tr><td>Fluorescent alignment target&amp;nbsp;</td><td>Thorlabs&amp;nbsp;</td><td>VRC2SM1</td><td></td></tr><tr><td>Laser safety goggles</td><td>Thorlabs&amp;nbsp;</td><td>LG3</td><td></td></tr><tr><td>ND filter(s)</td><td>Thorlabs&amp;nbsp;</td><td>FW1AND</td><td></td></tr><tr><td>Optical beam profiler</td><td>Thorlabs&amp;nbsp;</td><td>BP209-VIS</td><td></td></tr><tr><td>Post-mounted iris diaphragm</td><td>Thorlabs&amp;nbsp;</td><td>ID25</td><td>Quantity: 2</td></tr><tr><td>Shearing interferometer</td><td>Thorlabs&amp;nbsp;</td><td>SI100</td><td></td></tr><tr><td>XYZ translation stage, &amp;frac12;&amp;rdquo; travel&amp;nbsp;</td><td>Thorlabs&amp;nbsp;</td><td>T12XYZ</td><td></td></tr><tr><td>Laser</td><td>World Star Technologies&amp;nbsp;</td><td>TECGL-30</td><td>532 nm, 30 mW</td></tr></tbody></table>

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 ...

fluorescence-imaging-with-one-nanometer-accuracy-fiona

Research

JoVE Journal

Biology

17.6K 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

Compact Quantum Dots for Single-molecule Imaging

Single-molecule imaging is an important tool for understanding the mechanisms of biomolecular function and for visualizing the spatial and temporal heterogeneity of molecular behaviors that underlie cellular biology 1-4. To image an individual molecule of interest, it is typically conjugated to a fluorescent tag (dye, protein, bead, or quantum dot) and observed with epifluorescence or total internal reflection fluorescence (TIRF) microscopy. While dyes and fluorescent proteins have been the mainstay of fluorescence imaging for decades, their fluorescence is unstable under high photon fluxes necessary to observe individual molecules, yielding only a few seconds of observation before complete loss of signal. Latex beads and dye-labeled beads provide improved signal stability but at the expense of drastically larger hydrodynamic size, which can deleteriously alter the diffusion and behavior of the molecule under study.
Quantum dots (QDs) offer a balance between these two problematic regimes. These nanoparticles are composed of semiconductor materials and can be engineered with a hydrodynamically compact size with exceptional resistance to photodegradation 5. Thus in recent years QDs have been instrumental in enabling long-term observation of complex macromolecular behavior on the single molecule level. However these particles have still been found to exhibit impaired diffusion in crowded molecular environments such as the cellular cytoplasm and the neuronal synaptic cleft, where their sizes are still too large 4,6,7.
Recently we have engineered the cores and surface coatings of QDs for minimized hydrodynamic size, while balancing offsets to colloidal stability, photostability, brightness, and nonspecific binding that have hindered the utility of compact QDs in the past 8,9. The goal of this article is to demonstrate the synthesis, modification, and characterization of these optimized nanocrystals, composed of an alloyed HgxCd1-xSe core coated with an insulating CdyZn1-yS shell, further coated with a multidentate polymer ligand modified with short polyethylene glycol (PEG) chains (Figure 1). Compared with conventional CdSe nanocrystals, HgxCd1-xSe alloys offer greater quantum yields of fluorescence, fluorescence at red and near-infrared wavelengths for enhanced signal-to-noise in cells, and excitation at non-cytotoxic visible wavelengths. Multidentate polymer coatings bind to the nanocrystal surface in a closed and flat conformation to minimize hydrodynamic size, and PEG neutralizes the surface charge to minimize nonspecific binding to cells and biomolecules. The end result is a brightly fluorescent nanocrystal with emission between 550-800 nm and a total hydrodynamic size near 12 nm. This is in the same size range as many soluble globular proteins in cells, and substantially smaller than conventional PEGylated QDs (25-35 nm).

We describe the preparation of colloidal quantum dots with minimized hydrodynamic size for single-molecule fluorescence imaging. Compared to conventional quantum dots, these nanoparticles are similar in size to globular proteins and are optimized for single-molecule brightness, stability against photodegradation, and resistance to nonspecific binding to proteins and cells.

Single-molecule imaging is an important tool for  understanding the mechanisms of biomolecular function and for visualizing the  spatial and temporal ...

Engineering

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Developmental Biology

High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., kinetics, coexistence of different states and populations, transient interactions), which are typically hidden in ensemble measurements, such as those obtained with standard biochemical or microscopy methods. Thus, dynamic events, such as receptor-receptor interactions, can be followed in real time in a living cell with high spatiotemporal resolution. This protocol describes a method based on labeling with small and bright organic fluorophores and total internal reflection fluorescence (TIRF) microscopy to directly visualize single receptors on the surface of living cells. This approach allows one to precisely localize receptors, measure the size of receptor complexes, and capture dynamic events such as transient receptor-receptor interactions. The protocol provides a detailed description of how to perform a single-molecule experiment, including sample preparation, image acquisition and image analysis. As an example, the application of this method to analyze two G-protein-coupled receptors, i.e., &#946;2-adrenergic and &#947;-aminobutyric acid type B (GABAB) receptor, is reported. The protocol can be adapted to other membrane proteins and different cell models, transfection methods and labeling strategies.

This protocol describes how to use total internal reflection fluorescence microscopy to visualize and track single receptors on the surface of living cells and thereby analyze receptor lateral mobility, size of receptor complexes as well as to visualize transient receptor-receptor interactions. This protocol can be extended to other membrane proteins.

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., ...

Bioengineering

From Fast Fluorescence Imaging to Molecular Diffusion Law on Live Cell Membranes in a Commercial Microscope

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the regulation of many cellular functions. However, due to the fast dynamics and the tiny structures involved, a very high spatio-temporal resolution is required to catch the real behavior of molecules. Here we present the experimental protocol for studying the dynamics of fluorescently-labeled plasma-membrane proteins and lipids in live cells with high spatiotemporal resolution. Notably, this approach doesn&#8217;t need to track each molecule, but it calculates population behavior using all molecules in a given region of the membrane. The starting point is a fast imaging of a given region on the membrane. Afterwards, a complete spatio-temporal autocorrelation function is calculated correlating acquired images at increasing time delays, for example each 2, 3, n repetitions. It is possible to demonstrate that the width of the peak of the spatial autocorrelation function increases at increasing time delay as a function of particle movement due to diffusion. Therefore, fitting of the series of autocorrelation functions enables to extract the actual protein mean square displacement from imaging (iMSD), here presented in the form of apparent diffusivity vs average displacement. This yields a quantitative view of the average dynamics of single molecules with nanometer accuracy. By using a GFP-tagged variant of the Transferrin Receptor (TfR) and an ATTO488 labeled 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphoethanolamine (PPE) it is possible to observe the spatiotemporal regulation of protein and lipid diffusion on &#181;m-sized membrane regions in the micro-to-milli-second time range.

Spatial distribution and temporal dynamics of plasma membrane proteins and lipids is a hot topic in biology. Here this issue is addressed by a spatio-temporal image fluctuation analysis that provides conceptually the same physical quantities of single particle tracking, but it uses small molecular labels and standard microscopy setups.

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the ...

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular level. To date the shear complexity of biological systems has caused precise single-molecule experimentation to be far too demanding, instead focusing on studies of single systems using relatively crude bulk ensemble-average measurements. However, many important processes occur in the living cell at the level of just one or a few molecules; ensemble measurements generally mask the stochastic and heterogeneous nature of these events. Here, using advanced optical microscopy and analytical image analysis tools we demonstrate how to monitor proteins within a single living bacterial cell to a precision of single molecules and how we can observe dynamics within molecular complexes in functioning biological machines. The techniques are directly relevant physiologically. They are minimally-perturbative and non-invasive to the biological sample under study and are fully attuned for investigations in living material, features not readily available to other single-molecule approaches of biophysics. In addition, the biological specimens studied all produce fluorescently-tagged protein at levels which are almost identical to the unmodified cell strains ("genomic encoding"), as opposed to the more common but less ideal approach for generating significantly more protein than would occur naturally ('plasmid expression'). Thus, the actual biological samples which will be investigated are significantly closer to the natural organisms, and therefore the observations more relevant to real physiological processes.

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Here we demonstrate the protocols for performing single-molecule fluorescence microscopy on living bacterial cells to enable functional molecular complexes to be detected, tracked and quantified.

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular ...

Conducting Multiple Imaging Modes with One Fluorescence Microscope

Fluorescence microscopy is a powerful tool to detect biological molecules in situ and monitor their dynamics and interactions in real-time. In addition to conventional epi-fluorescence microscopy, various imaging techniques have been developed to achieve specific experimental goals. Some of the widely used techniques include single-molecule fluorescence resonance energy transfer (smFRET), which can report conformational changes and molecular interactions with angstrom resolution, and single-molecule detection-based super-resolution (SR) imaging, which can enhance the spatial resolution approximately ten&#160;to twentyfold compared to diffraction-limited microscopy. Here we present a customer-designed integrated system, which merges multiple imaging methods in one microscope, including conventional epi-fluorescent imaging, single-molecule detection-based SR imaging, and multi-color single-molecule detection, including smFRET imaging. Different imaging methods can be achieved easily and reproducibly by switching optical elements. This set-up is easy to adopt by any research laboratory in biological sciences with a need for routine and diverse imaging experiments at a reduced cost and space relative to building separate microscopes for individual purposes.

Here we present a practical guide of building an integrated microscopy system, which merges conventional epi-fluorescent imaging, single-molecule detection-based super-resolution imaging, and multi-color single-molecule detection, including single-molecule fluorescence resonance energy transfer imaging, into one set-up in a cost-efficient way.

Fluorescence microscopy is a powerful tool to detect biological molecules in situ and monitor their dynamics and interactions in real-time. In addition to ...

Three-dimensional Imaging of Bacterial Cells for Accurate Cellular Representations and Precise Protein Localization

The shape of a bacterium is important for its physiology. Many aspects of cell physiology such as cell motility, predation, and biofilm production can be affected by cell shape. Bacterial cells are three-dimensional (3D) objects, although they are rarely treated as such. Most microscopy techniques result in two-dimensional (2D) images leading to the loss of data pertaining to the actual 3D cell shape and localization of proteins. Certain shape parameters, such as Gaussian curvature (the product of the two principal curvatures), can only be measured in 3D because 2D images do not measure both principal curvatures. Additionally, not all cells lie flat when mounting and 2D imaging of curved cells may not accurately represent the shapes of these cells. Accurately measuring protein localization in 3D can help determine the spatial regulation and function of proteins. A forward convolution technique has been developed that uses the blurring function of the microscope to reconstruct 3D cell shapes and to accurately localize proteins. Here, a protocol for preparing and mounting samples for live cell imaging of bacteria in 3D both to reconstruct an accurate cell shape and to localize proteins is described. The method is based on simple sample preparation, fluorescent image acquisition, and MATLAB-based image processing. Many high-quality fluorescent microscopes can be simply modified to take these measurements. These cell reconstructions are computationally intensive and access to high-throughput computational resources is recommended, although not necessary. This method has been successfully applied to multiple bacterial species and mutants, fluorescent imaging modalities, and microscope manufacturers.

This protocol explains how to prepare and mount bacterial samples for live three-dimensional imaging and how to reconstruct the three-dimensional shape of E. coli from those images.

The shape of a bacterium is important for its physiology. Many aspects of cell physiology such as cell motility, predation, and biofilm production can be ...

Immunology and Infection

Super-resolution Imaging of the Bacterial Division Machinery

Bacterial cell division requires the coordinated assembly of more than ten essential proteins at midcell1,2. Central to this process is the formation of a ring-like suprastructure (Z-ring) by the FtsZ protein at the division plan3,4. The Z-ring consists of multiple single-stranded FtsZ protofilaments, and understanding the arrangement of the protofilaments inside the Z-ring will provide insight into the mechanism of Z-ring assembly and its function as a force generator5,6. This information has remained elusive due to current limitations in conventional fluorescence microscopy and electron microscopy. Conventional fluorescence microscopy is unable to provide a high-resolution image of the Z-ring due to the diffraction limit of light (~200 nm). Electron cryotomographic imaging has detected scattered FtsZ protofilaments in small C. crescentus cells7, but is difficult to apply to larger cells such as E. coli or B. subtilis. Here we describe the application of a super-resolution fluorescence microscopy method, Photoactivated Localization Microscopy (PALM), to quantitatively characterize the structural organization of the E. coli Z-ring8.
	PALM imaging offers both high spatial resolution (~35 nm) and specific labeling to enable unambiguous identification of target proteins. We labeled FtsZ with the photoactivatable fluorescent protein mEos2, which switches from green fluorescence (excitation = 488 nm) to red fluorescence (excitation = 561 nm) upon activation at 405 nm9. During a PALM experiment, single FtsZ-mEos2 molecules are stochastically activated and the corresponding centroid positions of the single molecules are determined with &lt;20 nm precision. A super-resolution image of the Z-ring is then reconstructed by superimposing the centroid positions of all detected FtsZ-mEos2 molecules.
	Using this method, we found that the Z-ring has a fixed width of ~100 nm and is composed of a loose bundle of FtsZ protofilaments that overlap with each other in three dimensions. These data provide a springboard for further investigations of the cell cycle dependent changes of the Z-ring10 and can be applied to other proteins of interest.

We describe a super-resolution imaging method to probe the structural organization of the bacterial FtsZ-ring, an essential apparatus for cell division. This method is based on quantitative analyses of photoactivated localization microscopy (PALM) images and can be applied to other bacterial cytoskeletal proteins.

Bacterial cell division requires the coordinated  assembly of more than ten essential proteins at midcell1,2. Central  to this process is the formation of ...

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 ...

Combining Single-molecule Manipulation and Imaging for the Study of Protein-DNA Interactions

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method offers the opportunity of investigating interactions occurring in solution (thus avoiding problems due to closeby surfaces as in other single molecule methods), controlling the DNA extension and tracking interaction dynamics as a function of both mechanical parameters and DNA sequence. The methods for establishing successful optical trapping and nanometer localization of single molecules are illustrated. We illustrate the experimental conditions allowing the study of interaction of lactose repressor (lacI), labeled with Atto532, with a DNA molecule containing specific target sequences (operators) for LacI binding. The method allows the observation of specific interactions at the operators, as well as one-dimensional diffusion of the protein during the process of target search. The method is broadly applicable to the study of protein-DNA interactions but also to molecular motors, where control of the tension applied to the partner track polymer (for example actin or microtubules) is desirable.

Here we describe the instrumentation and methods for detecting single fluorescently-labeled protein molecules interacting with a single DNA molecule suspended between two optically trapped microspheres.

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method ...

Fluorescence Imaging with One-nanometer Accuracy (FIONA)

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