This work was supported by Defense Advanced Research Projects Agency (DARPA) (HR00111720066) and National Science Foundation (NSF) (1805200). We thank Michael Serge in Andor Technology for generously loaning the sCMOS camera.

1. Setting up the microscope, lasers and alignment tools
<ol>
	<li>Before building the microscope, prepare all the necessary components including optics, optomechanics and electronics as listed in Table of Materials.</li>
	<li>Prepare a microscope body mainly composed of two parts: an objective holder with an RMS-threaded port and a piezo-stage mounted on an aluminum block (Figure 1A). 
	NOTE: The custom-made microscope body is used for the convenience and flexibility of instrumentation12. Any commercially available microscope body can be used for HIST microscopy.</li>
	<li>Combining multiple laser lines and coupling them to a single mode fiber
	<ol>
		<li>Install 405, 561, 638 nm lasers onto the optical table and combine the beams through a polarizing beam splitter and a long-pass dichroic mirror as shown in Figure 1B. Make sure that all laser beams pass through pinholes on the alignment tool. Insert half wave plates for power adjustment. 
		NOTE: Wear safety goggles for eye protection and use beam-blocks to absorb unwanted laser beams.</li>
		<li>Install a fiber coupling lens (f = 4.5 mm) and a fiber adapter held in z-axis translator with a cage system.</li>
		<li>Connect a multimode fiber (MMF, &#216; 62.5 &#181;m) to the fiber adapter. Adjust each pair of the steering mirror and the z-translator until the coupling efficiency of each laser is higher than 95%. The output beam has a near Gaussian-shaped profile with speckle patterns.</li>
		<li>Take away the multimode fiber and connect a single mode fiber (SMF). Similar to MMF, fine-tune and maximize the coupling efficiency of three lasers.</li>
	</ol>
	</li>
	<li>Assemble a collimated light source that will be used for beam alignment in the excitation and detection paths. This device is composed of a temporally coherent light source (561 nm) connected to SMF, fiber adapter, achromatic lens (f = 60 mm), iris and &#216;1&#34; tube spacer in a cage system (Figure 1C). Adjust the distance between the fiber adapter and the lens using a shearing interferometer to ensure the collimation.</li>
	<li>Prepare a beam alignment tool (Figure 1D). This is a pair of aluminum posts with pinholes at 2&#34; height from the surface of optical table, which allows for quick and precise beam alignment.</li>
	<li>Assemble a double pinhole system which consists of two &#216;1&#34; ground glass alignment disks at each end and two &#216;1&#34; lens tubes (the bottom one is slotted) as shown in Figure 1E.</li>
</ol>
2. Setting up the detection path
<ol>
	<li>Take out the objective and install the collimated light source. Adjust the knobs of the mirror (M1) beneath the objective holder so that the output beam from the microscope is roughly parallel to the optical table in height and aligned with threaded holes on the table. Refer to Figure 2 for detailed positions of each optical component.</li>
	<li>Insert a multiband dichroic mirror (DM) and reflect the beam by 90 degrees. Use the maximum size of the iris and make sure that the beam passes through the center of the dichroic mirror without clipping.</li>
	<li>Use the leaky beam passing through the dichroic mirror to guide the alignment of detection path. Place an sCMOS camera into the beam and make sure that the beam hits the center of the camera chip by using two mirrors (M2 and M3).</li>
	<li>Insert a tube lens (TL; f = 300 mm) approximately 300 mm away from the camera.</li>
	<li>Remove the collimated light source and adjust the relative distance between the tube lens and the camera until a pattern on the ceiling is clearly resolved by the camera.</li>
	<li>Insert a multi-band pass filter (BF) before the tube lens for multi-color fluorescence imaging.</li>
</ol>
3. Setting up the excitation path
<ol>
	<li>Reinstall the collimated light source on the objective holder. Place a fold-mirror (M4) to redirect the beam output from the microscope by 90 degrees. Adjust the knobs of the dichroic mirror and the fold-mirror iteratively until the beam passes through pinholes in the beam alignment tool.</li>
	<li>Detach the collimated light source and install it on the table, in which the beam points towards the microscope body. Align the beam using a beam alignment tool and a double pinhole system.</li>
	<li>Insert a lens L4 (f = 400 mm; &#216; = 2&#34;) to the optical path approximately 400 mm away from the objective holder. Install the objective lens and adjust the position of L4 along the optical axis until a perfect Airy disk pattern is formed on the ceiling. 
	NOTE: When inserting the lens, the beam position passing through the lens should be kept unchanged. The lens L4 has a SM2 thread that allows it to be attached/detached from the 60 mm SM2-threaded cage plate easily.</li>
	<li>Unscrew the objective and reinstall the collimated light source with an open iris. Trace down the beam output from the microscope with a business card. Mount a mirror M5 at the place where the size of beam is the smallest and approximately 400 mm away from L4, which is a conjugated image plane (cIP).</li>
	<li>Install a mirror M6 and reflect the beam by 90 degrees. Adjust M5 and M6 iteratively with the beam alignment tool.</li>
	<li>Insert a lens L3 (f = 150 mm) roughly 150 mm away from M5. Use a shearing interferometer to ensure the collimation of the output beam.</li>
	<li>Temporarily take away L4 and trace down the beam to find the focal position of L3. Put a single axis galvo mirror at this point, which is a conjugated back focal plane (cBFP). Supply 0 volts to the galvo mirror and rotate the holder of the galvo mirror so that it reflects the beam by 90 degrees.</li>
	<li>Place a fold-mirror M7. Properly place L2 (f = 100 mm) the same way as step 3.6.</li>
	<li>Remove the collimated light source from the object holder. Install a collimation lens L1 (f = 100 mm), fiber adapter and iris. Connect a single mode fiber to the adapter and send the beam through the imaging system.</li>
	<li>Reinsert L4 and fine tune the system until a perfect Airy disk pattern appears on the ceiling.</li>
</ol>
4. Setting up the cylindrical lenses
<ol>
	<li>Insert a cylindrical lens (CL1, f = 400 mm) after L1 and make sure that the cylindrical lens focuses the beam along the x-axis.</li>
	<li>Insert another cylindrical lens (CL2, f = 50 mm) to the beam path. Use a shearing interferometer to ensure the output beam is collimated. 
	NOTE: The distance between the two cylindrical lenses is 450 mm. The output beam has a compression ratio of 8 and an elongated oval shaped Airy disk pattern is formed on the ceiling.</li>
</ol>
5. Testing tile imaging
<ol>
	<li>Prepare 3D hydrogel sample. Mix 20 nm Crimson beads with a hydrogel solution, which consists of 12% acrylamide:bisacrylamide (29:1), 0.2% (v/v) TEMED and 0.2% (w/v) ammonium persulfate in 0.75x TAE buffer. Inject 50 &#181;L of the mixed solution into a flow chamber as described elsewhere11. After 10 min, the 3D hydrogel sample is ready to for imaging.</li>
	<li>Put the sample onto the sample holder. Turn on 638 nm laser and adjust the power to &#60;1 mW for sample excitation.</li>
	<li>Run the camera control software. In the camera acquisition-setting panel, select Internal in trigger mode and then click Take video for free running mode.</li>
	<li>Slightly adjust the position of the camera so that the image is located at the center of the camera when 0 volts is applied to the galvo mirror.</li>
	<li>Rotate the horizontal knob of the mirror M5 to achieve a highly inclined illumination. 
	NOTE: With the increase of the illumination angle, the hydrogel image becomes clearer than that of the Epi image as the beam gets thinner. However, the image keeps almost the same position.</li>
	<li>Record the tile image. Calculate the effective illumination width11. For example, Figure 3 shows the effective illumination width of 12 &#181;m.</li>
</ol>
6. HIST imaging
<ol>
	<li>Prepare a data acquisition board connected with a terminal block. Connect USER 1 BNC connector with P0.0 through an electrical wire. Use USER 1 as a digital output for the external triggering of sCMOS camera. Connect an analog output AO0 to a galvo mirror driver.</li>
	<li>Generate TTL pulse trains from P0.0 using a custom-made program (Figure 4A) and set the period time = 400 ms and t_ON = 2 ms. Check the generated pulses from USER 1 BNC terminal by a digital oscilloscope and then connect the BNC cable to the camera external trigger port. 
	NOTE: The control software used in this paper is available upon request. When imaging at different camera frame rates, the period time should be adjusted accordingly.</li>
	<li>Start sweeping a galvo mirror via the custom-made program. Adjust Vmin to -500 mV and Vmax to 500 mV for full FOV imaging. Note that under this operation, 3D hydrogel samples still show high background similar to Epi illumination.</li>
	<li>Changing the camera acquisition setting.
	<ol>
		<li>Select External in trigger mode and Down (Sequential) in LightScan Plus drop-down menu as shown in Figure 4B. 
		NOTE: In this setting, the camera does not take images unless a trigger signal is turned on.</li>
		<li>Click Scan Speed Control for window height and line exposure time control and set the values to be 180 rows and 28 ms, respectively. 
		NOTE: When a tile width (Weff) is 180 rows (12 &#181;m) and an integration time per line (Tint) is 28 ms, a delay time between lines (TD) is determined as TD = Tint/Weff = 0.156 ms. For imaging 2,048 x 2,048 pixels, the total acquisition time is 2,048 x TD + Tint = 346 ms, corresponding to ~2.9 fps.</li>
	</ol>
	</li>
	<li>Slightly adjust Vmax and Vmin to obtain clearer images.</li>
	<li>Obtain 3D stack images using the custom-made program by switching on 3D stack ON and specifying the number of stacks and the step size.</li>
</ol>

<ol>
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M., Zhou, R., Zhuang, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualizing+and+discovering+cellular+structures+with+super-resolution+microscopy.">Visualizing and discovering cellular structures with super-resolution microscopy.</a> Science. 361, 880-887 (2018).</li><li>Lerner, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Toward+dynamic+structural+biology:+Two+decades+of+single-molecule+Forster+resonance+energy+transfer.">Toward dynamic structural biology: Two decades of single-molecule Forster resonance energy transfer.</a> Science. 359, (2018).</li><li>Raj, A., van Oudenaarden, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-Molecule+Approaches+to+Stochastic+Gene+Expression.">Single-Molecule Approaches to Stochastic Gene Expression.</a> Annual Review of Biophysics. 38, 255-270 (2009).</li><li>Wilson, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Resolution+and+optical+sectioning+in+the+confocal+microscope.">Resolution and optical sectioning in the confocal microscope.</a> Journal of microscopy. 244, 113-121 (2011).</li><li>Sase, I., Miyata, H., Corrie, J. 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University of Central Florida has filed a patent application covering the work described in this paper.

There are two critical steps in this protocol. The first one is the proper placement of L4 in step 3.3, ensuring that the incident beam passes through the center of the lens and a perfect Airy disk pattern is formed on the ceiling. The position of L4 determines the placement of all the other optical components, including M5, L3, GM and L2. The second critical step is the synchronization process. To reject the out of focus background, active pixels whose effective detection width is equal to the tile width should be synchronized with the beam sweeping. Therefore, it is necessary to measure the effective illumination width of a tile beam (step 5.6) and set camera parameters accordingly in step 6.4.
When imaging with very large FOV, the presented method shows an increased background at one side compared to the other side. This is attributed to slightly altered angles of illumination at different imaging positions. Implementing a second galvo mirror instead of M5 alleviates this problem as demonstrated before by synchronously adjusting the position and the scanning angle11. Instead of off-the-shelf achromatic doublets, a telecentric scan lens will be also helpful. However, for imaging an area of &#60;8,080 &#181;m2, single galvo mirror sweeping was sufficient. HIST microscopy has a limit of the imaging depth, however, it is able to obtain a good SBR when imaging up to ~15 &#181;m with a 12 &#181;m tile beam and a NA 1.45 oil immersion objective lens11.
In this protocol, we used a beam compression ratio of 8 to make a tile beam. A thinner illumination can be used in HIST microscopy to achieve higher SBR, which might be powerful for single-molecule tissue imaging11. However, in this case, photobleaching effect should be considered by an increased excitation intensity while the current beam compression ratio showed reduced photobleaching in 3D imaging compared to Epi11. Compared to light-sheet microscopes with two orthogonally placed objectives, HIST microscopy is simple to implement and compatible with conventional sample preparations. The enhanced SBR and large FOV of HIST microscopy is suitable for studying the interactions and dynamics of single biomolecules in multiple cells and can be used further in super-resolution imaging and single-molecule tracking.

Single-molecule fluorescence imaging plays an important role in many biological studies that reveal ultrastructures, dynamics and the quantity of biomolecules1,2,3. However, it has been challenging to study single-molecules inside cells or tissues. While confocal microscopy provides high sectioning capability4, it is not suitable for single-molecule imaging due to severe photobleaching by the high excitation intensity or slow imaging speed. Widefield microscopy uses weaker illumination but suffers from a poor signal to background ratio (SBR)5. Light-sheet microscopy, on the other hand, could show good sectioning and low photobleaching6; however, the available numerical aperture (NA) is greatly limited by the requirement of orthogonally placed objectives7. Alternatively, it requires special illuminators and sample chambers8,9.
For these reasons, highly inclined and laminated optical sheet (HILO) microscopy has been widely used for 3D single-molecule imaging10. When an inclined beam encounters an interface of two media (glass and water, for example), the beam is refracted according to Snell&#39;s law. Importantly, the refracted beam gets thinner, and its thickness is described as dz = R/tan(&#952;) where R is the diameter of the inclined beam and &#952; is the refraction angle of the transmitted beam. This simple implementation results in a good sectioning capability. Nevertheless, this relation indicates that a thin illumination (i.e., high sectioning capability) requires a small R and/or a large &#952;. For example, when R = 20 &#181;m and &#952; = 72 degrees, one can obtain dz = 6.5 &#181;m. Since there is a practical limit to increasing the refraction angle in order to image deep inside cells and avoid total internal reflection, there is a strong coupling of the illumination diameter and the beam thickness. For this reason, HILO imaging shows a relatively small field-of-view (FOV) that greatly restricts its applications in multicellular imaging.
Recently, we have overcome this problem by highly inclined swept tile (HIST) microscopy where the FOV is decoupled from the beam thickness in a very simple way11. First, a beam elongated in one direction is generated via a pair of cylindrical lenses. This beam, termed as a tile, produces a thin illumination with dz ~ 4 &#181;m while its FOV is 130&#160;x 12 &#181;m2. Then, the tile is swept across the sample using a rotating galvo mirror. Meanwhile, the fluorescence image is recorded on a scientific complementary metal-oxide semiconductor (sCMOS) camera that filters efficiently out-of-focus background by operating in a rolling shutter mode which serves as tunable confocal slit detection. In this way, HIST microscopy enables single-molecule imaging with a larger field of view (~130&#160;x 130 &#181;m2) and a thinner illumination than HILO imaging. We applied this new imaging technique to detect RNA transcripts with a single probe in cells or with a few probes in mouse brain tissues, which has significant potential for studying gene expression and diseases. Unlike other approaches, HIST employs only a single high numerical aperture objective without an additional illuminator or remote detection objectives and is fully compatible with inverted microscopes. These advantages along with a large FOV and high contrast will make HIST microscopy a prominent tool in biology and medicine. We present detailed instructions regarding instrumentation of the HIST microscope, and how to test and calibrate its performance as below.

<table border="0" cellpadding="0" cellspacing="0" style="width:680px;" width="680">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">1&#34; Achromatic doublet</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">AC254-060-A-ML</td>
			<td style="width:225px;">Collimator&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">1&#34; Achromatic doublet</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">AC254-100-A-ML</td>
			<td style="width:225px;">L1,L2</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">1&#34; Achromatic doublet</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">AC254-300-A-ML</td>
			<td style="width:225px;">TL</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">1&#34; Broadband Dielectric Mirrors</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">BB1-E02-10</td>
			<td style="width:225px;">M1~M7</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">1&#34; Cylindrical Lenses</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">LJ1363RM-A</td>
			<td style="width:225px;">CL1</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">1&#34; Cylindrical Lenses</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">LJ1695RM-A</td>
			<td style="width:225px;">CL2</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">1&#34; square kinematic mount</td>
			<td style="width:88px;">Edmund Optics</td>
			<td style="width:151px;">58-857</td>
			<td style="width:225px;">For dichroic mirror mounting</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">1&#34; Threaded Cage Plate</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">CP02</td>
			<td style="width:225px;">For holding other lenses</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">2&#34; Achromatic doublet</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">AC508-150-A-ML</td>
			<td style="width:225px;">L3</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">2&#34; Achromatic doublet</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">AC508-400-A-ML</td>
			<td style="width:225px;">L4</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">2&#34; Threaded Cage Plate</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">LCP01</td>
			<td style="width:225px;">For holding L4</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">2&#34; Threaded Cage Plate</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">LCP01T</td>
			<td style="width:225px;">For holding L3</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">2% Bis Solution</td>
			<td style="width:88px;">Bio Rad</td>
			<td style="width:151px;">64085292</td>
			<td style="width:225px;">hydrogel component</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">20 nm fluorescent beads</td>
			<td style="width:88px;">Thermo Fisher</td>
			<td style="width:151px;">F8782</td>
			<td style="width:225px;">For testing imaging</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">30 mm Cage Right-Angle Kinematic Mirror Mount</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">KCB1</td>
			<td style="width:225px;">For objective &#38; camera mounting</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">30mm Cage System Iris</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">CP20S</td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">3-Axis NanoMax Stage</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">MAX311D</td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">40% Acrylamide Solution</td>
			<td style="width:88px;">Bio Rad</td>
			<td style="width:151px;">64148001</td>
			<td style="width:225px;">hydrogel component</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">405 nm laser</td>
			<td style="width:88px;">Cobolt</td>
			<td style="width:151px;">Cobolt 06-MLD</td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">50x TAE buffer</td>
			<td style="width:88px;">Bio-Rad</td>
			<td style="width:151px;">161-0743</td>
			<td style="width:225px;">hydrogel component</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">561 nm laser</td>
			<td style="width:88px;">Cobolt</td>
			<td style="width:151px;">Cobolt 06-DPL</td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">638 nm laser</td>
			<td style="width:88px;">Cobolt</td>
			<td style="width:151px;">Cobolt 06-MLD</td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Ammonium persulfate</td>
			<td style="width:88px;">Sigma</td>
			<td style="width:151px;">A3678-25G</td>
			<td style="width:225px;">hydrogel component</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Beam alignment tool</td>
			<td style="width:88px;">custom made</td>
			<td style="width:151px;"></td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">BNC terminal blocks</td>
			<td style="width:88px;">Natural Instruments</td>
			<td style="width:151px;">BNC-2110</td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">Cage plate with M9 x 0.5 internal threads</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">CP1TM09</td>
			<td style="width:225px;">For holding aspheric lens</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Cage System Rods</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">SR series</td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Cell culture &#38; smFISH</td>
			<td style="width:88px;"></td>
			<td style="width:151px;"></td>
			<td style="width:225px;">See a reference [11]</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Double side tape&#160;</td>
			<td style="width:88px;">Scotch</td>
			<td style="width:151px;">515182</td>
			<td style="width:225px;">Flow chamber</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Epoxy&#160;</td>
			<td style="width:88px;">Devcon</td>
			<td style="width:151px;">14250</td>
			<td style="width:225px;">Flow chamber</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Galvo mirror</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">GVS211</td>
			<td style="width:225px;">GM</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Galvo System Linear Power Supply</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">GPS011</td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Half wave plate</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">WPH10M-405/561/633</td>
			<td style="width:225px;">Power adjustment</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">long-pass dichroic mirror</td>
			<td style="width:88px;">Chroma</td>
			<td style="width:151px;">T550lpxr</td>
			<td style="width:225px;">For combining lasers</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Microscope slides</td>
			<td style="width:88px;">Fisherbrand</td>
			<td style="width:151px;">12549-3</td>
			<td style="width:225px;">Flow chamber</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">Mikroskopische Deckglaser</td>
			<td style="width:88px;">Hecht Assistent</td>
			<td style="width:151px;">990/5024</td>
			<td style="width:225px;">Flow chamber</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">Mounted Frosted Glass Alignment Disk</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">DG10-1500-H1-MD</td>
			<td style="width:225px;">For double pinhole system</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Mounted rochester aspheric lens</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">A230TM-A</td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">Multi-band dichroic mirror</td>
			<td style="width:88px;">Semrock</td>
			<td style="width:151px;">Di03-R405/488/561/635-t3</td>
			<td style="width:225px;">DM; 3 mm thickness</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">Multi-band filter</td>
			<td style="width:88px;">Semrock</td>
			<td style="width:151px;">FF01-446/523/600/677-25</td>
			<td style="width:225px;">BF</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Multimode fiber</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">M31L02</td>
			<td style="width:225px;">MMF</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">N,N,N&#39;,N&#39;-tetramethyl ethylenediamine</td>
			<td style="width:88px;">Sigma</td>
			<td style="width:151px;">T7024-25ML</td>
			<td style="width:225px;">hydrogel component</td>
		</tr>
		<tr height="35">
			<td height="35" style="height:35px;width:216px;">NI-DAQ board</td>
			<td style="width:88px;">Natural Instruments</td>
			<td style="width:151px;">PCI-6733</td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">&#216;1&#34; Kinematic Mirror Mount</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">KM100</td>
			<td style="width:225px;">For holding mirrors</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Objective lens</td>
			<td style="width:88px;">Olympus</td>
			<td style="width:151px;">PLANAPO N 60X</td>
			<td style="width:225px;">60X 1.45NA oil</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Pedestal Base Clamping Forks</td>
			<td style="width:88px;">Newport</td>
			<td style="width:151px;">9916</td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Pedestal Pillar Posts</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">RS1P8E</td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Piezo controller</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">BPC303</td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Polarized beam splitter</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">PBS251</td>
			<td style="width:225px;">For combining lasers</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">RMS-SM1 adapter</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">SM1A3TS</td>
			<td style="width:225px;">For objective lens</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Rod holder</td>
			<td style="width:88px;">custom made</td>
			<td style="width:151px;"></td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Rotation cage mount&#160;</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">RSP1/CRM1/CRM1P</td>
			<td style="width:225px;">For HWP &#38; cylindrical lens mounting</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">sCMOS camera</td>
			<td style="width:88px;">Andor</td>
			<td style="width:151px;">Zyla-4.2P-CL10</td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Shearing interferometer</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">SI100</td>
			<td style="width:225px;">Beam collimation test</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Single mode fiber</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">P5-405BPM-FC-2</td>
			<td style="width:225px;">SMF</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">SM1 Lens Tubes</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">SM1S25</td>
			<td style="width:225px;">For double pinhole system</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">SM1 Slotted Lens Tube</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">SM1L30C</td>
			<td style="width:225px;">For double pinhole system</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Stage mount</td>
			<td style="width:88px;">custom made</td>
			<td style="width:151px;"></td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">threaded fiber adapter</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">SM1FC</td>
			<td style="width:225px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Z-Axis Translation Mount</td>
			<td style="width:88px;">Thorlabs</td>
			<td style="width:151px;">SM1Z</td>
			<td style="width:225px;">Fiber coupling</td>
		</tr>
	</tbody>
</table>

a guide to build a highly inclined swept tile microscope for extended field-of-view single-molecule imaging

Single-molecule imaging has greatly advanced our understanding of molecular mechanisms in biological studies. However, it has been challenging to obtain large field-of-view, high-contrast images in thick cells and tissues. Here, we introduce highly inclined swept tile (HIST) microscopy that overcomes this problem. A pair of cylindrical lenses was implemented to generate an elongated excitation beam that was scanned over a large imaging area via a fast galvo mirror. A 4f configuration was used to position optical components. A scientific complementary metal-oxide semiconductor camera detected the fluorescence signal and blocked the out-of-focus background with a dynamic confocal slit synchronized with the beam sweeping. We present a step-by-step instruction on building the HIST microscope with all basic components.

As an example, single-stranded DNA labeled with Atto647N was imaged with an excitation wavelength of 638 nm in a 3D hydrogel. DNA was anchored to the hydrogel network via an acrydite moiety during gel polymerization. The images were taken at 5 &#181;m above the surface as shown in Figure 5a. The HIST image showed much less background compared to the Epi image, from which the signal to background ratio was calculated to be 1.9 &#177; 0.7 for the HIST image while most of the single molecule spots could barely be detected by Epi.
Single-molecule RNA fluorescence in situ hybridization (smFISH) was performed with 4 FISH probes. Figure 5b displays smFISH images of EEF2 (eukaryotic translation elongation factor 2) labeled with AlexaFluor 647 on A549 cells in an imaging buffer (refer to our previous work regarding the sample preparation11). A maximum intensity projection was performed on 20 z-stacks corresponding to 5 &#181;m thickness. The HIST image showed not only much improved SBR but also more uniform illumination compared to Epi image. For Epi imaging, the exposure time was 400 ms while for HIST imaging the integration time per line was 32 ms, both of which had the same illumination power of 7.5 mW measured before the objective. The imaging speeds of Epi and HIST were 2.5 fps.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59360/59360fig1.jpg" /> 
Figure 1. Microscope body, lasers and alignment tools. (A) Objective and sample holder. (B) Photo of laser systems. LP, long-pass dichroic mirror; &#955;/2, half-wave plates; PBS, polarizing beamsplitter. (C) Collimated light source. (D) Beam alignment tool with two insertable pinholes. (E) Double pinhole system. <a href="https://www.jove.com/files/ftp_upload/59360/59360fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59360/59360fig2.jpg" /> 
Figure 2. Detailed setup for highly inclined swept tile (HIST) microscopy. Photo (A) and schematic (B) of HIST microscope system. BF, multi-band pass filter; CL1-2, cylindrical lenses; DM, dichroic mirror; GM, galvo mirror; BF, band-pass filter; M1-7, mirrors; L1-4, lenses; SMF, single mode fiber; TL, tube lens; cIP, conjugated image plane; cBFP, conjugated back focal plane. <a href="https://www.jove.com/files/ftp_upload/59360/59360fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59360/59360fig3.jpg" /> 
Figure 3. Tile illumination with a compression ratio of 8. (A) Fluorescence image of 20 nm beads in a 3D hydrogel. Scale bar, 20 &#181;m. (B) Standard deviation projection along the y direction of A, smoothed by 10 data points. The red arrow indicates an effective illumination width of 12 &#181;m. <a href="https://www.jove.com/files/ftp_upload/59360/59360fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59360/59360fig4v2.jpg" /> 
Figure 4. Control and imaging software front panels. (A) A custom-made LabView program synchronously controls the scanning of the galvo mirror, the starting acquisition of sCMOS camera and the movement of the piezo stage. (B) Camera acquisition setting control panel. <a href="https://www.jove.com/files/ftp_upload/59360/59360fig4v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/59360/59360fig5.jpg" /> 
Figure 5. (A) Images of Atto647N-labeled DNA in a 3D hydrogel with Epi and HIST illumination. (B) smFISH images of EEF2 using 4 FISH probes on A549 cells by Epi and HIST microscopy. DAPI stain is shown in blue. Scale bars, 20 &#181;m. <a href="https://www.jove.com/files/ftp_upload/59360/59360fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Guide to Build a Highly Inclined Swept Tile Microscope for Extended Field-of-view Single-molecule Imaging at JoVE.com

A Guide to Build a Highly Inclined Swept Tile Microscope for Extended Field-of-view Single-molecule Imaging

A detailed instruction is described on how to build a highly inclined swept tile (HIST) microscope and its usage for single-molecule imaging.

Single-molecule imaging has greatly advanced our understanding of molecular mechanisms in biological studies. However, it has been challenging to obtain ...

a-guide-to-build-highly-inclined-swept-tile-microscope-for-extended

Research

JoVE Journal

Bioengineering

17.0K Views.  University of Central Florida. A detailed instruction is described on how to build a highly inclined swept tile (HIST) microscope and its usage for single-molecule imaging.

Video: A Guide to Build a Highly Inclined Swept Tile Microscope for Extended Field-of-view Single-molecule Imaging

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

Investigating Receptor-ligand Systems of the Cellulosome with AFM-based Single-molecule Force Spectroscopy

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the enzymes onto the noncatalytic scaffold protein is directed by interactions among a family of related receptor-ligand pairs comprising interacting cohesin and dockerin modules. The extremely strong binding between cohesin and dockerin modules results in dissociation constants in the low picomolar to nanomolar range, which may hamper accurate off-rate measurements with conventional bulk methods. Single-molecule force spectroscopy (SMFS) with the atomic force microscope measures the response of individual biomolecules to force, and in contrast to other single-molecule manipulation methods (i.e.&#160;optical tweezers), is optimal for studying high-affinity receptor-ligand interactions because of its ability to probe the high-force regime (&#62;120 pN). Here we present our complete protocol for studying cellulosomal protein assemblies at the single-molecule level. Using a protein topology derived from the native cellulosome, we worked with enzyme-dockerin and carbohydrate binding module-cohesin (CBM-cohesin) fusion proteins, each with an accessible free thiol group at an engineered cysteine residue. We present our site-specific surface immobilization protocol, along with our measurement and data analysis procedure for obtaining detailed binding parameters for the high-affinity complex. We demonstrate how to quantify single subdomain unfolding forces, complex rupture forces, kinetic off-rates, and potential widths of the binding well. The successful application of these methods in characterizing the cohesin-dockerin interaction responsible for assembly of multidomain cellulolytic complexes is further described.

Cellulosomes are multienzyme complexes designed for digesting cellulose. AFM-based SMFS was used to study the mechanical properties and folding configuration of cellulosome-associated protein assemblies. We present a complete workflow for protein immobilization, data acquisition, and data analysis to study the interactions of individual receptor-ligand complexes involved in cellulosome assembly.

Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the ...

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., ...

3D Orbital Tracking in a Modified Two-photon Microscope: An Application to the Tracking of Intracellular Vesicles

The objective of this video protocol is to discuss how to perform and analyze a three-dimensional fluorescent orbital particle tracking experiment using a modified two-photon microscope1. As opposed to conventional approaches (raster scan or wide field based on a stack of frames), the 3D orbital tracking allows to localize and follow with a high spatial (10 nm accuracy) and temporal resolution (50 Hz frequency response) the 3D displacement of a moving fluorescent particle on length-scales of hundreds of microns2. The method is based on a feedback algorithm that controls the hardware of a two-photon laser scanning microscope in order to perform a circular orbit around the object to be tracked: the feedback mechanism will maintain the fluorescent object in the center by controlling the displacement of the scanning beam3-5. To demonstrate the advantages of this technique, we followed a fast moving organelle, the lysosome, within a living cell6,7. Cells were plated according to standard protocols, and stained using a commercially lysosome dye. We discuss briefly the hardware configuration and in more detail the control software, to perform a 3D orbital tracking experiment inside living cells. We discuss in detail the parameters required in order to control the scanning microscope and enable the motion of the beam in a closed orbit around the particle. We conclude by demonstrating how this method can be effectively used to track the fast motion of a labeled lysosome along microtubules in 3D within a live cell. Lysosomes can move with speeds in the range of 0.4-0.5 &#181;m/sec, typically displaying a directed motion along the microtubule network8.

In this video protocol we track - at high speed and in three dimensions - fluorescently labeled lysosomes within living cells, using the orbital tracking method in a modified two-photon microscope.

The objective of this video protocol is to discuss how to perform and analyze a three-dimensional fluorescent orbital particle tracking experiment using a ...

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 ...

Microfluidic Genipin Deposition Technique for Extended Culture of Micropatterned Vascular Muscular Thin Films

The chronic nature of vascular disease progression requires the development of experimental techniques that simulate physiologic and pathologic vascular behaviors on disease-relevant time scales. Previously, microcontact printing has been used to fabricate two-dimensional functional arterial mimics through patterning of extracellular matrix protein as guidance cues for tissue organization. Vascular muscular thin films utilized these mimics to assess functional contractility. However, the microcontact printing fabrication technique used typically incorporates hydrophobic PDMS substrates. As the tissue turns over the underlying extracellular matrix, new proteins must undergo a conformational change or denaturing in order to expose hydrophobic amino acid residues to the hydrophobic PDMS surfaces for attachment, resulting in altered matrix protein bioactivity, delamination, and death of the tissues.
Here, we present a microfluidic deposition technique for patterning of the crosslinker compound genipin. Genipin serves as an intermediary between patterned tissues and PDMS substrates, allowing cells to deposit newly-synthesized extracellular matrix protein onto a more hydrophilic surface and remain attached to the PDMS substrates. We also show that extracellular matrix proteins can be patterned directly onto deposited genipin, allowing dictation of engineered tissue structure. Tissues fabricated with this technique show high fidelity in both structural alignment and contractile function of vascular smooth muscle tissue in a vascular muscular thin film model. This technique can be extended using other cell types and provides the framework for future study of chronic tissue- and organ-level functionality.

We present a method for microfluidic deposition of patterned genipin and fibronectin on PDMS substrates, allowing extended viability of vascular smooth muscle cell-dense tissues. This tissue fabrication method is combined with previous vascular muscular thin film technology to measure vascular contractility over disease-relevant time courses.

The chronic nature of vascular disease progression requires the development of experimental techniques that simulate physiologic and pathologic vascular ...

Single Molecule Fluorescence Microscopy on Planar Supported Bilayers

In the course of a single decade single molecule microscopy has changed from being a secluded domain shared merely by physicists with a strong background in optics and laser physics to a discipline that is now enjoying vivid attention by life-scientists of all venues 1. This is because single molecule imaging has the unique potential to reveal protein behavior in situ in living cells and uncover cellular organization with unprecedented resolution below the diffraction limit of visible light 2. Glass-supported planar lipid bilayers (SLBs) are a powerful tool to bring cells otherwise growing in suspension in close enough proximity to the glass slide so that they can be readily imaged in noise-reduced Total Internal Reflection illumination mode 3,4. They are very useful to study the protein dynamics in plasma membrane-associated events as diverse as cell-cell contact formation, endocytosis, exocytosis and immune recognition. Simple procedures are presented how to generate highly mobile protein-functionalized SLBs in a reproducible manner, how to determine protein mobility within and how to measure protein densities with the use of single molecule detection. It is shown how to construct a cost-efficient single molecule microscopy system with TIRF illumination capabilities and how to operate it in the experiment.

Preparation of protein-functionalized planar glass-supported lipid bilayers, determination of protein mobility within and measurement of protein densities is shown here. A roadmap to building a noise-reduced Total Internal Reflection microscope is outlined, which allows visualizing single bilayer-resident fluorochromes with high spatiotemporal resolution.

In the course of a single decade single molecule microscopy has changed from being a secluded domain shared merely by physicists with a strong background ...

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 ...

Using Nanoplasmon-Enhanced Scattering and Low-Magnification Microscope Imaging to Quantify Tumor-Derived Exosomes

Infected or malignant cells frequently secrete more exosomes, leading to elevated levels of disease-associated exosomes in the circulation. These exosomes have the potential to serve as biomarkers for disease diagnosis and to monitor disease progression and treatment response. However, most exosome analysis procedures require exosome isolation and purification steps, which are usually time-consuming and labor-intensive, and thus of limited utility in clinical settings. This report describes a rapid procedure to analyze specific biomarkers on the outer membrane of exosomes without requiring separate isolation and purification steps. In this method, exosomes are captured on the surface of a slide by exosome-specific antibodies and then hybridized with nanoparticle-conjugated antibody probes specific to a disease. After hybridization, the abundance of the target exosome population is determined by analyzing low-magnification dark-field microscope (LMDFM) images of the bound nanoparticles. This approach can be easily adopted for research and clinical use to analyze membrane-associated exosome biomarkers linked to disease.

Clinical translation of exosome-derived biomarkers for diseased and malignant cells is hindered by the lack of rapid and accurate quantification methods. This report describes the use of low-magnification dark-field microscope images to quantify specific exosome subtypes in small volume serum or plasma samples.

Infected or malignant cells frequently secrete more exosomes, leading to elevated levels of disease-associated exosomes in the circulation. These exosomes ...

Fabrication of Zero Mode Waveguides for High Concentration Single Molecule Microscopy

In single molecule fluorescence enzymology, background fluorescence from labeled substrates in solution often limits fluorophore concentration to pico- to nanomolar ranges, several orders of magnitude less than many physiological ligand concentrations. Optical nanostructures called zero mode waveguides (ZMWs), which are 100&#8722;200 nm in diameter apertures fabricated in a thin conducting metal such as aluminum or gold, allow imaging of individual molecules at micromolar concentrations of fluorophores by confining visible light excitation to zeptoliter effective volumes. However, the need for expensive and specialized nanofabrication equipment has precluded the widespread use of ZMWs. Typically, nanostructures such as ZMWs are obtained by direct writing using electron beam lithography, which is sequential and slow. Here, colloidal, or nanosphere, lithography is used as an alternative strategy to create nanometer-scale masks for waveguide fabrication. This report describes the approach in detail, with practical considerations for each phase. The method allows thousands of aluminum or gold ZMWs to be made in parallel, with final waveguide diameters and depths of 100&#8722;200 nm. Only common lab equipment and a thermal evaporator for metal deposition are required. By making ZMWs more accessible to the biochemical community, this method can facilitate the study of molecular processes at cellular concentrations and rates.

Described here is a nanosphere lithography method for parallel fabrication of zero mode waveguides, which are arrays of nanoapertures in a metal-clad glass microscopy coverslip for single molecule imaging at nano- to micromolar concentrations of fluorophores. The method takes advantage of colloidal crystal self-assembly to create a waveguide template.

In single molecule fluorescence enzymology, background fluorescence from labeled substrates in solution often limits fluorophore concentration to pico- to ...