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><li>Sigal, Y. M., Zhou, R., Zhuang, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Visualizing+and+discovering+cellular+structures+with+super-resolution+microscopy%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">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/pubmed?term=((Toward+dynamic+structural+biology%3A+Two+decades+of+single-molecule+Forster+resonance+energy+transfer%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">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/pubmed?term=((Single-Molecule+Approaches+to+Stochastic+Gene+Expression%5BTitle%5D)+AND+%22Annual+Review+of+Biophysics%22%5BJournal%5D)">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/pubmed?term=((Resolution+and+optical+sectioning+in+the+confocal+microscope%5BTitle%5D)+AND+%22Journal+of+microscopy%22%5BJournal%5D)">Resolution and optical sectioning in the confocal microscope.</a> Journal of microscopy. 244, 113-121 (2011).</li>
<li>Sase, I., Miyata, H., Corrie, J. E., Craik, J. S., Kinosita, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Real+time+imaging+of+single+fluorophores+on+moving+actin+with+an+epifluorescence+microscope%5BTitle%5D)+AND+%22Biophysical+Journal%22%5BJournal%5D)">Real time imaging of single fluorophores on moving actin with an epifluorescence microscope.</a> Biophysical Journal. 69, 323-328 (1995).</li>
<li>Huisken, J., Swoger, J., Del Bene, F., Wittbrodt, J., Stelzer, E. H. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Optical+Sectioning+Deep+Inside+Live+Embryos+by+Selective+Plane+Illumination+Microscopy%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">Optical Sectioning Deep Inside Live Embryos by Selective Plane Illumination Microscopy.</a> Science. 305, 1007-1009 (2004).</li>
<li>Cella Zanacchi, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Live-cell+3D+super-resolution+imaging+in+thick+biological+samples%5BTitle%5D)+AND+%22Nature+Methods%22%5BJournal%5D)">Live-cell 3D super-resolution imaging in thick biological samples.</a> Nature Methods. 8, 1047(2011).</li>
<li>Galland, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((3D+high-+and+super-resolution+imaging+using+single-objective+SPIM%5BTitle%5D)+AND+%22Nature+Methods%22%5BJournal%5D)">3D high- and super-resolution imaging using single-objective SPIM.</a> Nature Methods. 12, 641(2015).</li>
<li>Gustavsson, A. K., Petrov, P. N., Lee, M. Y., Shechtman, Y., Moerner, W. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((3D+single-molecule+super-resolution+microscopy+with+a+tilted+light+sheet%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">3D single-molecule super-resolution microscopy with a tilted light sheet.</a> Nature Communications. 9, 123(2018).</li>
<li>Tokunaga, M., Imamoto, N., Sakata-Sogawa, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Highly+inclined+thin+illumination+enables+clear+single-molecule+imaging+in+cells%5BTitle%5D)+AND+%22Nature+Methods%22%5BJournal%5D)">Highly inclined thin illumination enables clear single-molecule imaging in cells.</a> Nature Methods. 5, 159-161 (2008).</li>
<li>Tang, J., Han, K. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Extended+field-of-view+single-molecule+imaging+by+highly+inclined+swept+illumination%5BTitle%5D)+AND+%22Optica%22%5BJournal%5D)">Extended field-of-view single-molecule imaging by highly inclined swept illumination.</a> Optica. 5, 1063-1069 (2018).</li>
<li>Han, K. Y., Kim, S. K., Eggeling, C., Hell, S. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Metastable+Dark+States+Enable+Ground+State+Depletion+Microscopy+of+Nitrogen+Vacancy+Centers+in+Diamond+with+Diffraction-Unlimited+Resolution%5BTitle%5D)+AND+%22Nano+Letters%22%5BJournal%5D)">Metastable Dark States Enable Ground State Depletion Microscopy of Nitrogen Vacancy Centers in Diamond with Diffraction-Unlimited Resolution.</a> Nano Letters. 10, 3199-3203 (2010).</li>
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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><tbody><tr><td>1&quot; Achromatic doublet</td><td>Thorlabs</td><td>AC254-060-A-ML</td><td>Collimator&amp;nbsp;</td></tr><tr><td>1&quot; Achromatic doublet</td><td>Thorlabs</td><td>AC254-100-A-ML</td><td>L1,L2</td></tr><tr><td>1&quot; Achromatic doublet</td><td>Thorlabs</td><td>AC254-300-A-ML</td><td>TL</td></tr><tr><td>1&quot; Broadband Dielectric Mirrors</td><td>Thorlabs</td><td>BB1-E02-10</td><td>M1~M7</td></tr><tr><td>1&quot; Cylindrical Lenses</td><td>Thorlabs</td><td>LJ1363RM-A</td><td>CL1</td></tr><tr><td>1&quot; Cylindrical Lenses</td><td>Thorlabs</td><td>LJ1695RM-A</td><td>CL2</td></tr><tr><td>1&quot; square kinematic mount</td><td>Edmund Optics</td><td>58-857</td><td>For dichroic mirror mounting</td></tr><tr><td>1&quot; Threaded Cage Plate</td><td>Thorlabs</td><td>CP02</td><td>For holding other lenses</td></tr><tr><td>2&quot; Achromatic doublet</td><td>Thorlabs</td><td>AC508-150-A-ML</td><td>L3</td></tr><tr><td>2&quot; Achromatic doublet</td><td>Thorlabs</td><td>AC508-400-A-ML</td><td>L4</td></tr><tr><td>2&quot; Threaded Cage Plate</td><td>Thorlabs</td><td>LCP01</td><td>For holding L4</td></tr><tr><td>2&quot; Threaded Cage Plate</td><td>Thorlabs</td><td>LCP01T</td><td>For holding L3</td></tr><tr><td>2% Bis Solution</td><td>Bio Rad</td><td>64085292</td><td>hydrogel component</td></tr><tr><td>20 nm fluorescent beads</td><td>Thermo Fisher</td><td>F8782</td><td>For testing imaging</td></tr><tr><td>30 mm Cage Right-Angle Kinematic Mirror Mount</td><td>Thorlabs</td><td>KCB1</td><td>For objective &amp;amp; camera mounting</td></tr><tr><td>30mm Cage System Iris</td><td>Thorlabs</td><td>CP20S</td><td></td></tr><tr><td>3-Axis NanoMax Stage</td><td>Thorlabs</td><td>MAX311D</td><td></td></tr><tr><td>40% Acrylamide Solution</td><td>Bio Rad</td><td>64148001</td><td>hydrogel component</td></tr><tr><td>405 nm laser</td><td>Cobolt</td><td>Cobolt 06-MLD</td><td></td></tr><tr><td>50x TAE buffer</td><td>Bio-Rad</td><td>161-0743</td><td>hydrogel component</td></tr><tr><td>561 nm laser</td><td>Cobolt</td><td>Cobolt 06-DPL</td><td></td></tr><tr><td>638 nm laser</td><td>Cobolt</td><td>Cobolt 06-MLD</td><td></td></tr><tr><td>Ammonium persulfate</td><td>Sigma</td><td>A3678-25G</td><td>hydrogel component</td></tr><tr><td>Beam alignment tool</td><td>custom made</td><td></td><td></td></tr><tr><td>BNC terminal blocks</td><td>Natural Instruments</td><td>BNC-2110</td><td></td></tr><tr><td>Cage plate with M9 x 0.5 internal threads</td><td>Thorlabs</td><td>CP1TM09</td><td>For holding aspheric lens</td></tr><tr><td>Cage System Rods</td><td>Thorlabs</td><td>SR series</td><td></td></tr><tr><td>Cell culture &amp;amp; smFISH</td><td></td><td></td><td>See a reference [11]</td></tr><tr><td>Double side tape&amp;nbsp;</td><td>Scotch</td><td>515182</td><td>Flow chamber</td></tr><tr><td>Epoxy&amp;nbsp;</td><td>Devcon</td><td>14250</td><td>Flow chamber</td></tr><tr><td>Galvo mirror</td><td>Thorlabs</td><td>GVS211</td><td>GM</td></tr><tr><td>Galvo System Linear Power Supply</td><td>Thorlabs</td><td>GPS011</td><td></td></tr><tr><td>Half wave plate</td><td>Thorlabs</td><td>WPH10M-405/561/633</td><td>Power adjustment</td></tr><tr><td>long-pass dichroic mirror</td><td>Chroma</td><td>T550lpxr</td><td>For combining lasers</td></tr><tr><td>Microscope slides</td><td>Fisherbrand</td><td>12549-3</td><td>Flow chamber</td></tr><tr><td>Mikroskopische Deckglaser</td><td>Hecht Assistent</td><td>990/5024</td><td>Flow chamber</td></tr><tr><td>Mounted Frosted Glass Alignment Disk</td><td>Thorlabs</td><td>DG10-1500-H1-MD</td><td>For double pinhole system</td></tr><tr><td>Mounted rochester aspheric lens</td><td>Thorlabs</td><td>A230TM-A</td><td></td></tr><tr><td>Multi-band dichroic mirror</td><td>Semrock</td><td>Di03-R405/488/561/635-t3</td><td>DM; 3 mm thickness</td></tr><tr><td>Multi-band filter</td><td>Semrock</td><td>FF01-446/523/600/677-25</td><td>BF</td></tr><tr><td>Multimode fiber</td><td>Thorlabs</td><td>M31L02</td><td>MMF</td></tr><tr><td>N,N,N&#039;,N&#039;-tetramethyl ethylenediamine</td><td>Sigma</td><td>T7024-25ML</td><td>hydrogel component</td></tr><tr><td>NI-DAQ board</td><td>Natural Instruments</td><td>PCI-6733</td><td></td></tr><tr><td>&amp;Oslash;1&quot; Kinematic Mirror Mount</td><td>Thorlabs</td><td>KM100</td><td>For holding mirrors</td></tr><tr><td>Objective lens</td><td>Olympus</td><td>PLANAPO N 60X</td><td>60X 1.45NA oil</td></tr><tr><td>Pedestal Base Clamping Forks</td><td>Newport</td><td>9916</td><td></td></tr><tr><td>Pedestal Pillar Posts</td><td>Thorlabs</td><td>RS1P8E</td><td></td></tr><tr><td>Piezo controller</td><td>Thorlabs</td><td>BPC303</td><td></td></tr><tr><td>Polarized beam splitter</td><td>Thorlabs</td><td>PBS251</td><td>For combining lasers</td></tr><tr><td>RMS-SM1 adapter</td><td>Thorlabs</td><td>SM1A3TS</td><td>For objective lens</td></tr><tr><td>Rod holder</td><td>custom made</td><td></td><td></td></tr><tr><td>Rotation cage mount&amp;nbsp;</td><td>Thorlabs</td><td>RSP1/CRM1/CRM1P</td><td>For HWP &amp;amp; cylindrical lens mounting</td></tr><tr><td>sCMOS camera</td><td>Andor</td><td>Zyla-4.2P-CL10</td><td></td></tr><tr><td>Shearing interferometer</td><td>Thorlabs</td><td>SI100</td><td>Beam collimation test</td></tr><tr><td>Single mode fiber</td><td>Thorlabs</td><td>P5-405BPM-FC-2</td><td>SMF</td></tr><tr><td>SM1 Lens Tubes</td><td>Thorlabs</td><td>SM1S25</td><td>For double pinhole system</td></tr><tr><td>SM1 Slotted Lens Tube</td><td>Thorlabs</td><td>SM1L30C</td><td>For double pinhole system</td></tr><tr><td>Stage mount</td><td>custom made</td><td></td><td></td></tr><tr><td>threaded fiber adapter</td><td>Thorlabs</td><td>SM1FC</td><td></td></tr><tr><td>Z-Axis Translation Mount</td><td>Thorlabs</td><td>SM1Z</td><td>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

Universidad Científica del Sur

Research

JoVE Journal

Bioengineering

17.4K 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

Single Plane Illumination Module and Micro-capillary Approach for a Wide-field Microscope

A module for light sheet or single plane illumination microscopy (SPIM) is described which is easily adapted to an inverted wide-field microscope and optimized for 3-dimensional cell cultures, e.g., multi-cellular tumor spheroids (MCTS). The SPIM excitation module shapes and deflects the light such that the sample is illuminated by a light sheet perpendicular to the detection path of the microscope. The system is characterized by use of a rectangular capillary for holding (and in an advanced version also by a micro-capillary approach for rotating) the samples, by synchronous adjustment of the illuminating light sheet and the objective lens used for fluorescence detection as well as by adaptation of a microfluidic system for application of fluorescent dyes, pharmaceutical agents or drugs in small quantities. A protocol for working with this system is given, and some technical details are reported. Representative results include (1) measurements of the uptake of a cytostatic drug (doxorubicin) and its partial conversion to a degradation product, (2) redox measurements by use of a genetically encoded glutathione sensor upon addition of an oxidizing agent, and (3) initiation and labeling of cell necrosis upon inhibition of the mitochondrial respiratory chain. Differences and advantages of the present SPIM module in comparison with existing systems are discussed.

A module for single plane illumination microscopy (SPIM) is described which is easily adapted to an inverted wide-field microscope and optimized for 3-dimensional cell cultures. The sample is located within a rectangular capillary, and via a microfluidic system fluorescent dyes, pharmaceutical agents or drugs can be applied in small quantities.

A module for light sheet or single plane illumination microscopy (SPIM) is described which is easily adapted to an inverted wide-field microscope and ...

Engineering

Single-Molecule Tracking Microscopy - A Tool for Determining the Diffusive States of Cytosolic Molecules

Single-molecule localization microscopy probes the position and motions of individual molecules in living cells with tens of nanometer spatial and millisecond temporal resolution. These capabilities make single-molecule localization microscopy ideally suited to study molecular level biological functions in physiologically relevant environments. Here, we demonstrate an integrated protocol for both acquisition and processing/analysis of single-molecule tracking data to extract the different diffusive states a protein of interest may exhibit. This information can be used to quantify molecular complex formation in living cells. We provide a detailed description of a camera-based 3D single-molecule localization experiment, as well as the subsequent data processing steps that yield the trajectories of individual molecules. These trajectories are then analyzed using a numerical analysis framework to extract the prevalent diffusive states of the fluorescently labeled molecules and the relative abundance of these states. The analysis framework is based on stochastic simulations of intracellular Brownian diffusion trajectories that are spatially confined by an arbitrary cell geometry. Based on the simulated trajectories, raw single-molecule images are generated and analyzed in the same way as experimental images. In this way, experimental precision and accuracy limitations, which are difficult to calibrate experimentally, are explicitly incorporated into the analysis workflow. The diffusion coefficient and relative population fractions of the prevalent diffusive states are determined by fitting the distributions of experimental values using linear combinations of simulated distributions. We demonstrate the utility of our protocol by resolving the diffusive states of a protein that exhibits different diffusive states upon forming homo- and hetero-oligomeric complexes in the cytosol of a bacterial pathogen.

3D single-molecule localization microscopy is utilized to probe the spatial positions and motion trajectories of fluorescently labeled proteins in living bacterial cells. The experimental and data analysis protocol described herein determines the prevalent diffusive behaviors of cytosolic proteins based on pooled single-molecule trajectories.

Single-molecule localization microscopy probes the position and motions of individual molecules in living cells with tens of nanometer spatial and ...

Biochemistry

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 ...

High-Throughput Total Internal Reflection Fluorescence and Direct Stochastic Optical Reconstruction Microscopy Using a Photonic Chip

Total internal reflection fluorescence (TIRF) is commonly used in single molecule localization based super-resolution microscopy as it gives enhanced contrast due to optical sectioning. The conventional approach is to use high numerical aperture microscope TIRF objectives for both excitation and collection, severely limiting the field of view and throughput. We present a novel approach to generating TIRF excitation for imaging with optical waveguides, called chip-based nanoscopy. The aim of this protocol is to demonstrate how chip-based imaging is performed in an already built setup. The main advantage of chip-based nanoscopy is that the excitation and collection pathways are decoupled. Imaging can then be done with a low magnification lens, resulting in large field of view TIRF images, at the price of a small reduction in resolution. Liver sinusoidal endothelial cells (LSECs) were imaged using direct stochastic optical reconstruction microscopy (dSTORM), showing a resolution comparable to traditional super-resolution microscopes. In addition, we demonstrate the high-throughput capabilities by imaging a 500 &#181;m x 500 &#181;m region with a low magnification lens, providing a resolution of 76 nm. Through its compact character, chip-based imaging can be retrofitted into most common microscopes and can be combined with other on-chip optical techniques, such as on-chip sensing, spectroscopy, optical trapping, etc. The technique is thus ideally suited for high throughput 2D super-resolution imaging, but also offers great opportunities for multi-modal analysis.

Chip-based super-resolution optical microscopy is a novel approach to fluorescence microscopy and offers advantages in cost effectiveness and throughput. Here, the protocols for chip preparation and imaging are shown for TIRF microscopy and localization-based super-resolution microscopy.

Total internal reflection fluorescence (TIRF) is commonly used in single molecule localization based super-resolution microscopy as it gives enhanced ...

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 ...

Optical Scatter Microscopy Based on Two-Dimensional Gabor Filters

We demonstrate a microscopic instrument that can measure subcellular texture arising from organelle morphology and organization within unstained living cells. The proposed instrument extends the sensitivity of label-free optical microscopy to nanoscale changes in organelle size and shape and can be used to accelerate the study of the structure-function relationship pertaining to organelle dynamics underlying fundamental biological processes, such as programmed cell death or cellular differentiation. The microscope can be easily implemented on existing microscopy platforms, and can therefore be disseminated to individual laboratories, where scientists can implement and use the proposed methods with unrestricted access.
The proposed technique is able to characterize subcellular structure by observing the cell through two-dimensional optical Gabor filters. These filters can be tuned to sense with nanoscale (10's of nm) sensitivity, specific morphological attributes pertaining to the size and orientation of non-spherical subcellular organelles. While based on contrast generated by elastic scattering, the technique does not rely on a detailed inverse scattering model or on Mie theory to extract morphometric measurements. This technique is therefore applicable to non-spherical organelles for which a precise theoretical scatter description is not easily given, and provides distinctive morphometric parameters that can be obtained within unstained living cells to assess their function. The technique is advantageous compared with digital image processing in that it operates directly on the object's field transform rather than the discretized object's intensity. It does not rely on high image sampling rates and can therefore be used to rapidly screen morphological activity within hundreds of cells at a time, thus greatly facilitating the study of organelle structure beyond individual organelle segmentation and reconstruction by fluorescence confocal microscopy of highly magnified digital images of limited fields of view.
In this demonstration we show data from a marine diatom to illustrate the methodology. We also show preliminary data collected from living cells to give an idea of how the method may be applied in a relevant biological context.

We demonstrate a dark-field microscopy method based on Gabor-like filtering to measure subcellular dynamics within single living cells. The technique is sensitive to alterations in the structure of organelles, such as mitochondrial fragmentation.

We demonstrate a microscopic instrument that can measure subcellular texture arising from organelle morphology and organization within unstained living ...

Biology

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 ...

Application of High-speed Super-resolution SPEED Microscopy in Live Primary Cilium

The primary cilium is a microtubule-based protrusion on the surface of many eukaryotic cells and contains a unique complement of proteins that function critically in cell motility and signaling. Since cilia are incapable of synthesizing their own protein, nearly 200 unique ciliary proteins need to be trafficked between the cytosol and primary cilia. However, it is still a technical challenge to map three-dimensional (3D) locations of transport pathways for these proteins in live primary cilia due to the limitations of currently existing techniques. To conquer the challenge, recently we have developed and employed a high-speed virtual 3D super-resolution microscopy, termed single-point edge-excitation sub-diffraction (SPEED) microscopy, to determine the 3D spatial location of transport pathways for both cytosolic and membrane proteins in primary cilia of live cells. In this article, we will demonstrate the detailed setup of SPEED microscopy, the preparation of cells expressing fluorescence-protein-labeled ciliary proteins, the real-time single-molecule tracking of individual proteins in live cilium and the achievement of 3D spatial probability density maps of transport routes for ciliary proteins.

Recently we mapped the three-dimensional (3D) spatial locations of transport routes for various proteins translocating inside primary cilia in live cells. Here this paper details the experimental setup, the process of biological samples and the data analyses for the 3D super-resolution fluorescence imaging approach newly applied in live primary cilia.

The primary cilium is a microtubule-based protrusion on the surface of many eukaryotic cells and contains a unique complement of proteins that function ...

Imaging Intermediate Filaments and Microtubules with 2-dimensional Direct Stochastic Optical Reconstruction Microscopy

The cytoskeleton, composed of actin microfilaments, microtubules, and intermediate filaments (IF), plays a key role in the control of cell shape, polarity, and motility. The organization of the actin and microtubule networks has been extensively studied but that of IFs is not yet fully characterized. IFs have an average diameter of 10 nm and form a network extending throughout the cell cytoplasm. They are physically associated with actin and microtubules through molecular motors and cytoskeletal linkers. This tight association is at the heart of the regulatory mechanisms that ensure the coordinated regulation of the three cytoskeletal networks required for most cell functions. It is therefore crucial to visualize IFs alone and also together with each of the other cytoskeletal networks. However, IF networks are extremely dense in most cell types, especially in glial cells, which makes its resolution very difficult to achieve with standard fluorescence microscopy (lateral resolution of ~250 nm). Direct STochastic Optical Reconstruction Microscopy (dSTORM) is a technique allowing a gain in lateral resolution of one order of magnitude. Here, we show that lateral dSTORM resolution is sufficient to resolve the dense organization of the IF networks and, in particular, of IF bundles surrounding microtubules. Such tight association is likely to participate in the coordinated regulation of these two networks and may, explain how vimentin IFs template and stabilize microtubule organization as well as could influence microtubule dependent vesicular trafficking. More generally, we show how the observation of two cytoskeletal components with dual-color dSTORM technique brings new insight into their mutual interaction.

The overall goal of this methodology is to give the optimal experimental conditions from sample preparation to image acquisition and reconstruction in order to perform 2D dual color dSTORM images of microtubules and intermediate filaments in fixed cells

The cytoskeleton, composed of actin microfilaments, microtubules, and intermediate filaments (IF), plays a key role in the control of cell shape, ...

Accelerating MIBI Run Setup Using TSAI for Tissue Microarray Imaging

Rapid Setup of Tissue Microarray and Tiled Area Imaging on the Multiplexed Ion Beam Imaging Microscope Using the Tile/SED/Array Interface

Multiplexed ion beam imaging (MIBI) is a next-generation mass spectrometry-based microscopy technique&#160;that generates 40+ plex images of protein expression in histologic tissues, enabling detailed dissection of cellular phenotypes and histoarchitectural organization. A key bottleneck in&#160;operation occurs when users select the physical locations on the tissue for imaging. As the scale and complexity of MIBI experiments have increased, the manufacturer-provided interface and third-party tools have become increasingly unwieldy for imaging large tissue microarrays and tiled tissue areas. Thus, a web-based, interactive, what-you-see-is-what-you-get (WYSIWYG) graphical interface layer - the tile/SED/array Interface (TSAI) - was developed for users to set imaging locations using familiar and intuitive mouse gestures such as drag-and-drop, click-and-drag, and polygon drawing. Written according to web standards already built into modern web browsers, it requires no installation of external programs, extensions, or compilers. Of interest to the hundreds of current MIBI users, this interface dramatically simplifies and accelerates the setup of large, complex MIBI runs.

Author Spotlight: Introducing the Tile/SED/Array Interface for Rapid Field of View Positioning in Tissue Imaging

Multiplexed ion beam imaging (MIBI) is often used to image tissue microarrays and tiled, contiguous tissue areas, but current software for setting up these experiments is cumbersome. The tile/SED/array interface is an intuitive, interactive graphical tool developed to dramatically simplify and accelerate MIBI run setup.

Multiplexed ion beam imaging (MIBI) is a next-generation mass spectrometry-based microscopy technique&#160;that generates 40+ plex images of protein ...