This work was supported by grants from the Leverhulme Trust, the Engineering and Physical Sciences Research Council [EP/H018301/1, EP/G037221/1]; Alzheimer Research UK [ARUK-EG2012A-1]; Wellcome Trust [089703/Z/09/Z] and Medical Research Council [MR/K015850/1, MR/K02292X/1]. We thank E. Avezov and M. Lu for transfection of the LifeAct-GFP and cytosolic-GFP cells respectively, and W. Chen for preparation of the HEK293 culture. We also thank K. O&#39;Holleran for assistance with the design of the microscope, and L. Shao and R. Heintzmann for useful discussions and suggestions.

1. Arranging and Aligning the Excitation Path
<ol>
	<li>Mark the positions of the components on the optical table (see Figure 1 for an overview of the optical setup). Separate the objective, lenses L3, L4, L5, and the SLM each by the sum of their respective focal lengths such that the SLM surface will be relayed onto the focal plane of the objective.</li>
	<li>Insert multi-edge dichroic mirror DM4 into the filter cube turret of the microscope frame.</li>
	<li>Insert the second dichroic mirror DM3 into a 1&#34; square kinetic mirror mount, and position it one focal length away from the condenser lens L5. 
	Note: This excitation path design incorporates two identical dichroic mirrors DM3 and DM4 which are taken from the same production batch to ensure identical optical properties. The dichroic mirror (DM4) is positioned such that the s- and p- axes are switched compared to the dichroic located in the microscope (DM3) thereby cancelling any polarization ellipticity introduced by its birefringence (Figure 1). This compensation works equally well for each illumination wavelength. This step is essential for maintaining high modulation contrast.</li>
	<li>Before inserting any lenses into the excitation path, accurately define the optical axis for the system.
	<ol>
		<li>Remove the objective lens (OB) from the turret and instead screw in an alignment tool. This consists of a 500 mm long optical cage system with two alignment disks at both ends.</li>
		<li>Use the dichroic mirror DM3 and a temporary alignment mirror positioned at the approximate later location of the SLM to steer a collimated reference beam from Laser 1 through the center of the holes in the two alignment disks. Direct the beam from Laser 1 to the temporary mirror as depicted in Figure 1 using three mirrors and dichroic mirror DM2. The temporary mirror at the SLM position must be close to perpendicular to the optical axis. 
		Note: Use Laser 1 as the reference beam, as the other lasers can be subsequently aligned once the excitation path is in place.</li>
		<li>Remove the alignment tool once the coarse optical axis has been determined.Insert an iris into the beam path before it enters the microscopy body and center it on the beam. Attach a piece of white card with a small hole centered on the iris.Reinsert the objective lens (OB). 
		Note: The beam leaving the objective will now be highly divergent, but there will be a very weak reflection from the back surface of the lens that will be visible on the white card. All lenses, even if they are anti-reflection coated, will have weak back reflections that can be used to ensure coaxial alignment. If the beam is exactly perpendicular to the lens then the back reflection will go back through the center of the iris</li>
		<li>Make iterative angular adjustments to the two mirrors (DM3 and alignment mirror at the SLM position) to center the back reflection on the card with the incoming beam. Temporarily remove the objective lens (OB) and mark the laser spot on the ceiling to create a reference position.</li>
		<li>Insert a pair of irises at the height of the reference beam along the threaded holes of the table. The beam should be parallel to the surface of the optical table. The optical axis is now defined.</li>
	</ol>
	</li>
	<li>Insert the condenser lens (L5) roughly one focal length away from the objective. Mount this lens on a linear translation stage set to translate along the direction of the reference beam.</li>
	<li>Adjust the condenser lens position and angle such that the beam leaving the objective is collimated and hits the reference spot on the ceiling. Check that the lens is perpendicular to the beam by again checking the back reflection with the iris and white card. Remove the objective lens (OB) and insert the second lens of the image relay telescope (L4). 
	Note: Ensuring proper collimation and non-deflection of the beam is made easier when there is an even number of lenses in the beam path.</li>
	<li>Adjust the position and angle of this lens using a linear translation stage to maintain collimation and to ensure the reference beam still hits the marked spot on the ceiling.</li>
	<li>Replace the objective lens (OB) and insert the first lens of the telescope (L3). Adjust the position and angle of this lens to ensure collimation and non-deflection, as described in previous steps.</li>
	<li>Mount the SLM chip on a gimbal mount which provides rotation without translation about the center of the chip surface. 
	Note: The specific mounting design depends on the SLM used. If the SLM is supplied without a mount, it should be fixed to a custom machined aluminum plate which is then attached to a lens gimbal mount.</li>
	<li>With the lenses aligned, insert the SLM in place of the mirror. Adjust the position of the SLM such that the reference beam is located at the center of the SLM chip, and adjust the angle such that the beam passes through the two relay lenses (L3 and L4). Check that the reference beam is still centered on the marked spot.</li>
	<li>Expand and collimate the reference beam using a Keplerian beam expander.
	<ol>
		<li>Mount the two lenses (L1 and L2) in a cage system for ease of adjustment.</li>
		<li>Centre the cage system on the reference beam by removing the lenses and replacing them with irises.</li>
		<li>Insert the two lenses and adjust the axial position of L2 to collimate the expanded beam using a shearing interferometer. L2 should be one focal length away from the surface of the SLM.</li>
		<li>Check that the expanded beam is still collimated after the two relay lenses L3 and L4. Use the shearing interferometer just after DM3 to check for collimation.</li>
	</ol>
	</li>
	<li>Once the excitation path has been aligned for a single wavelength, couple the other two lasers into the beam path. Steer each beam through two irises centered on the excitation path using the beam combining dichroic mirrors (DM1 and DM2).</li>
</ol>
2. Alignment of Polarization Rotator
<ol>
	<li>Mount the LCVR with its fast axis at 45&#176; to the incident polarization.</li>
	<li>Fine tune polarization angle of the beam incident to the LCVR using an achromatic half wave plate (HWP) by inserting the HWP and the LCVR between crossed polarizers. Rotate the HWP to minimize the transmitted power. 
	Note: In order to act as a variable polarization rotator, the fast axis of the liquid crystal retarder (LCVR) must be precisely aligned at 45&#176; to the incident vertical beam polarization. The LCVR is physically mounted at 45&#176; but this is only a coarse alignment. The HWP is used to ensure perfect 45&#176; alignment of the incident polarization with respect to the LCVR fast axis. The quarter wave plate (QWP) converts the tilted elliptical polarization induced by the LCVR back to linear polarization at an angle controlled by the applied voltage24.</li>
	<li>Insert the QWP after the LCVR and rotate it to align its slow axis to the incoming polarization by minimizing the transmitted power between crossed polarizers.</li>
</ol>
3. Alignment of the Emission Path
<ol>
	<li>Coarsely position the camera using a stage micrometer slide and transmitted light.
	<ol>
		<li>Focus on the reticle using the microscope oculars and fix the objective lens at this position.</li>
		<li>Roughly center the camera and move the camera position to bring the image of the reticle into focus by observing the image on screen. 
		Note: If an external filter wheel is used then the filter cube will not contain an emission filter, therefore the oculars must not be used when lasers are switched on.</li>
	</ol>
	</li>
	<li>Finely adjust the camera position using a fluorescent bead sample.
	<ol>
		<li>Prepare a monolayer of fluorescent beads by spreading a drop of 100 nm multicolor beads on a #1.5 coverglass. Leave to dry to adsorb the beads to the coverglass and then re-immerse in water.</li>
		<li>Place the bead sample onto the objective with immersion oil. Finely adjust the position of the camera such that the fluorescent bead layer is in focus. Do not adjust the objective lens position once the focus has been found. 
		Note: As the SLM must be in a plane conjugate to the sample plane, the position of the SLM, relay lenses, and objective must be fixed. To adjust the focus, move the sample axially instead of the objective using a piezo z-stage.</li>
	</ol>
	</li>
	<li>Generate the appropriate SIM binary grating patterns as bitmap files.
	<ol>
		<li>For 2D/TIRF-SIM, generate a series of 9 binary grating images: 3 pattern orientations each with 3 equally spaced phase shifts. Generate these numerically (using MATLAB for example) from a rotated 2D sinusoid with a phase offset applied, then thresholding to produce a binary image. See Supplemental Code Files for example code.</li>
		<li>For alignment purposes, also generate grating patterns that have been windowed by a small circular aperture for each of the 3 orientations, as shown in Figure 2. The windowed alignment gratings do not need to be externally triggered but can be manually switched by the user via the SLM&#39;s software. 
		Note: See references for a discussion of the optimal rotation angles and an example of grating pattern generation code 16,20.</li>
	</ol>
	</li>
	<li>Upload the bitmap images to the SLM using the manufacturer&#39;s software (for example MetroCon).
	<ol>
		<li>Load the SLM control software and click &#34;Connect&#34;.</li>
		<li>In the &#34;Repertoire&#34; tab, click &#34;Load&#34; to open the repertoire file and check the number of Running Orders contained in the file. In the example repertoire file given there are five Running Orders.</li>
		<li>Click &#34;Send to Board&#34; to upload the repertoire file to the SLM.</li>
		<li>Wait for the bitmap images to upload and for the device to automatically reboot. 
		Note: An example repertoire file, which contains grating bitmap images and a file defining the order, is included as a Supplemental Code File. The &#34;.repz&#34; file may be opened using ZIP file archiver software.</li>
	</ol>
	</li>
	<li>Display a windowed alignment grating on the SLM for the first orientation (for example 0&#176;).
	<ol>
		<li>In the SLM control software, select the &#34;Status&#34; tab, enter the number of the Running Order (in the case of the example file, this is Running Order &#34;1&#34;).</li>
		<li>Click &#34;Select&#34; to change the Running Order to the alignment grating. 
		Note: This will illuminate a small circular region in the sample plane. If the SLM surface is correctly conjugated to the sample plane then the edges of this region will be sharply in focus. The grating pattern will produce multiple diffraction orders at the focus of L3: the zero order reflection from the reflective backplane of the SLM, the -1 and +1 orders corresponding to the grating, and also weaker higher orders that arise from diffraction of internal elements specific to the SLM device (e.g. reflections of the internal wirings of the SLM pixels and irregularities at the pixel edges). All but the -1 and +1 orders must be filtered out.</li>
	</ol>
	</li>
	<li>Insert a spatial mask (SM) mounted in an x,y stage into the beam path at the focal position of L3, and translate its position with respect to the optical axis such that only the desired first orders are passed. Directly after the spatial filter, only two circular beams will be visible. 
	Note: The spatial mask is fabricated by punching 6 holes into aluminum foil using a needle. The holes should be large enough to pass the first order beams for all laser wavelengths. A detailed analysis of the spatial mask is given in reference20.</li>
	<li>Display the next orientation of the alignment grating (60&#176;, running order 2) and again ensure that only the first orders are let through the spatial mask, adjusting its position if required.</li>
	<li>Repeat for the final orientation (120&#176;, running order 3).</li>
	<li>Check the image of the fluorescent bead layer on the camera. If the two circular beams are not overlapping as depicted in Figure 2 then reposition the sample plane by iteratively adjusting the objective lens and camera position.</li>
	<li>Adjust the objective position to overlap the two beams which will bring the image out of focus. Reposition the camera to bring the image back into focus and fine tune the objective in case two circles are still visible. Repeat this process until the two beams overlap and a single circular region is in focus.</li>
	<li>Once the position of the sample plane has been set, keep the objective position fixed.</li>
	<li>To confirm TIRF illumination, image a solution of fluorescent dye, for example for a 488 nm excitation wavelength, use a solution of 10 &#181;M rhodamine 6G.
	<ol>
		<li>Bring the dye sample into focus. If the two beams are incident at the correct TIRF angle then single molecules will be visible without high background, and the edges of the circular aperture will be in focus. See Figure 2B-D for examples of aligned and misaligned TIRF beams.</li>
		<li>Display each orientation of the windowed gratings in turn and ensure that all three orientations provide TIRF illumination and that the two beams overlap at the sample plane. Fine adjustments to the position of the beams can be made by adjusting dichroic mirror DM3. 
		Note: Although different wavelengths are focused at slightly different positions due to axial chromatic aberration, this is not critical and may be corrected by applying a constant z-offset to the sample position prior to excitation with the second wavelength.</li>
	</ol>
	</li>
</ol>
4. System Synchronization and Calibration
<ol>
	<li>Place the bead monolayer sample on the objective and bring into focus.</li>
	<li>Program the SLM using its control software to display each of the 3 phase shift images in turn, for the first pattern orientation (0&#176;).
	<ol>
		<li>Using the SLM control software, switch to Running Order 4 of the example repertoire.</li>
		<li>Configure the camera using its acquisition software (for example HCImage) to output two signals: one positive and one negative TTL trigger signal during the global exposure period. In the camera software, under &#34;Advanced Camera Properties&#34;, set Output Trigger Kind 1 and 2 to &#34;Exposure&#34;, and Output Trigger Polarity 1 and 2 to &#34;Positive&#34; and &#34;Negative&#34; respectively.</li>
		<li>Connect Output 1 and 2 of the camera to the &#34;Trigger&#34; and &#34;Finish&#34; inputs of the SLM respectively, using coaxial cables. The SLM is now synchronized to the camera.</li>
	</ol>
	</li>
	<li>Acquire a series of 3 images.
	<ol>
		<li>In the &#34;Sequence&#34; pane, select &#34;Hard Disk Record&#34; as the scan type, and set the frame count to 3.</li>
		<li>Click &#34;Start&#34; to acquire 3 frames. The SLM pattern will change upon each exposure. The fluorescent beads in the image will appear to blink on and off between each of the 3 images. The amount of blinking is a read out of the modulation contrast of the sinusoidal illumination pattern.</li>
	</ol>
	</li>
	<li>Rotate the polarization of the excitation laser with the LCVR using custom software in order to achieve azimuthal polarization and therefore the highest modulation contrast for the given pattern orientation.
	<ol>
		<li>Load the LCVR calibration software.</li>
		<li>Enter 0 and 8 for the Minimum and Maximum Voltage respectively.</li>
		<li>Click &#34;Sweep LCVR Voltage&#34; to rotate the polarization. 
		Note: The LCVR retardance is a function of temperature and can drift day-to-day even with temperature control. In this step, optimal azimuthal polarization is found empirically by sweeping the applied voltage between its minimum and maximum voltage which has the effect of rotating the polarization incident at the sample. The modulation contrast is calculated for each voltage25 and the voltage that achieves peak contrast is used in the following steps.</li>
		<li>Wait for the calibration process to complete, and note down the measured voltage.</li>
	</ol>
	</li>
	<li>Repeat this calibration process for the remaining two pattern orientations (60&#176; and 120&#176;) and each of the excitation wavelengths.</li>
	<li>Synchronize the camera exposure with the LCVR, lasers, emission filter wheel and piezo z-stage26. To accomplish this, use a high speed data acquisition (DAQ) board as the master clock source for the system, and use the SLM&#39;s LED Enable output signal to modulate the lasers (see Figure 3B). 
	Note: The specific implementation is dependent on the components used but the use of a high speed DAQ board for digital trigger synchronization and control of the LCVR using an analog voltage, controlled via software, is recommended. The control software used in this protocol is available upon request.</li>
	<li>Due to axial chromatic aberration, for each wavelength, also apply a z-offset to the sample stage.
	<ol>
		<li>Determine the offset experimentally by focusing on a multicolor bead monolayer sample at the first wavelength (e.g. 488 nm) then switching to the second (e.g. 640 nm). The beads will now be out of focus.</li>
		<li>Refocus the beads and measure the change in z position that was needed. This offset can then be applied to the piezo z-stage every time the excitation wavelength is changed.</li>
	</ol>
	</li>
	<li>Using the SLM control software, switch the SLM Running Order to the full series of 9 binary grating images required for TIRF-SIM. This is Running Order 0 in the example repertoire.</li>
	<li>Using the camera control software, acquire 9 images of the bead sample.
	<ol>
		<li>In the &#34;Sequence&#34; pane of the camera software, select &#34;Hard Disk Record&#34; as the scan type, and change the frame count to 9.</li>
		<li>Click &#34;Start&#34; to acquire images.</li>
		<li>Save the acquired images as TIFF files by selecting &#34;TIFF&#34; as the image type in the &#34;Save Buffered Images&#34; window, and clicking OK.</li>
	</ol>
	</li>
	<li>Reconstruct a super-resolution image from the raw TIFF images using commercial or custom software to validate the improvement in resolution over standard TIRF. 
	Note: For our microscope we use custom reconstruction code developed both in-house and by Dr. Lin Shao27.</li>
</ol>

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Lett. 37 (22), 4603 (2012).</li><li>Brankner, S. Z., Hobson, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Synchronization and Triggering with the ORCA-Flash4.0 Scientific CMOS Camera. , (2013).</li><li>Gustafsson, M. G. L., 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=Three-dimensional+resolution+doubling+in+wide-field+fluorescence+microscopy+by+structured+illumination.">Three-dimensional resolution doubling in wide-field fluorescence microscopy by structured illumination.</a> Biophys. J. 94 (12), 4957-4970 (2008).</li><li>Wicker, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Non-iterative+determination+of+pattern+phase+in+structured+illumination+microscopy+using+auto-correlations+in+Fourier+space.">Non-iterative determination of pattern phase in structured illumination microscopy using auto-correlations in Fourier space.</a> Opt. Express. 21 (21), 24692 (2013).</li><li>Boulanger, J., Pustelnik, N., Condat, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Non-smooth+convex+optimization+for+an+efficient+reconstruction+in+structured+illumination+microscopy.">Non-smooth convex optimization for an efficient reconstruction in structured illumination microscopy.</a> 2014 IEEE 11th Int. Symp. Biomed. Imaging. 3 (1), 995-998 (2014).</li><li>Str&#246;hl, F., Kaminski, C. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+joint+Richardson-Lucy+deconvolution+algorithm+for+the+reconstruction+of+multifocal+structured+illumination+microscopy+data.">A joint Richardson-Lucy deconvolution algorithm for the reconstruction of multifocal structured illumination microscopy data.</a> Methods Appl. Fluoresc. 3 (1), 014002 (2015).</li><li>Mudry, 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=Structured+illumination+microscopy+using+unknown+speckle+patterns.">Structured illumination microscopy using unknown speckle patterns.</a> Nat. Photonics. 6 (5), 312-315 (2012).</li><li>Ayuk, R., 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=Structured+illumination+fluorescence+microscopy+with+distorted+excitations+using+a+filtered+blind-SIM+algorithm.">Structured illumination fluorescence microscopy with distorted excitations using a filtered blind-SIM algorithm.</a> Opt. Lett. 38 (22), 4723 (2013).</li><li>Ball, G., 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=SIMcheck:+a+Toolbox+for+Successful+Super-resolution+Structured+Illumination+Microscopy.">SIMcheck: a Toolbox for Successful Super-resolution Structured Illumination Microscopy.</a> Sci. Rep. 5, 15915 (2015).</li><li>York, A. G., 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=Resolution+doubling+in+live,+multicellular+organisms+via+multifocal+structured+illumination+microscopy.">Resolution doubling in live, multicellular organisms via multifocal structured illumination microscopy.</a> Nat. Methods. 9 (7), 749-754 (2012).</li><li>Li, D., 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=Extended-resolution+structured+illumination+imaging+of+endocytic+and+cytoskeletal+dynamics.">Extended-resolution structured illumination imaging of endocytic and cytoskeletal dynamics.</a> Science. 349 (6251), (2015).</li><li>York, A. G., 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=Resolution+doubling+in+live,+multicellular+organisms+via+multifocal+structured+illumination+microscopy.">Resolution doubling in live, multicellular organisms via multifocal structured illumination microscopy.</a> Nat. Methods. 9 (7), 749-754 (2012).</li></ol>

The authors have nothing to disclose.

Custom-built TIRF-SIM systems such as the setup detailed in this protocol are capable of multicolor super-resolution imaging at high speed compared to commercially available microscopes. The inherent advantage of SIM as a super-resolution technique is that the temporal resolution is not limited by the photophysics of the fluorophore, compared to other methods such as single molecule localization microscopy (SMLM) or point scanning methods such as stimulated emission depletion microscopy (STED). Unlike these other techniques, SIM does not require photoswitchable or depletable fluorophores so multicolor imaging is straightforward. Non TIRF-SIM systems, such as optical sectioning SIM and multifocal SIM can usually achieve resolution improvements of 1.7 times or less in practice as opposed to the factor of 2 improvement reported here, and commercial systems are also often slower and less flexible than the system presented in this protocol.
The two main difficulties in implementing this technique are firstly the necessity for precise positioning of the six SIM beams within the TIR zone of the objective&#39;s back aperture, which requires a laborious and time consuming optical alignment procedure. Secondly, to produce high pattern contrast at the sample, polarization rotation is essential. For low NA 2D-SIM systems, polarization rotation can be avoided by careful choice of the linear polarization orientation, but this becomes impossible for TIRF-SIM25. For high-speed multicolor imaging, electro-optical polarization control is necessary and this increases the complexity and expense of the system.
Limitations of the technique
TIRF-SIM, like conventional TIRF, is naturally limited to observation of biological structures and processes located at the basal cell membrane that can be illuminated by the 150-200 nm penetration depth of the evanescent field. While SIM is often quoted as being less photodamaging to cells than either STED or SMLM, lateral resolution doubling does still increase the required number of photons by at least 4-fold5 compared to conventional TIRF microscopy. For imaging at high frame rates with short exposures times, this photon increase necessitates use of increased illumination intensities. While any fluorophore may be used for SIM imaging of fixed or slow moving samples, high brightness fluorescent proteins or next generation synthetic dyes with enhanced photostability are recommended for live cell imaging.
Although this implementation is capable of imaging a single color at SIM frame rates in excess of 20 Hz, multicolor imaging in the presented system is limited by the switching time of the motorized emission filter wheel. Due to the large size of the sCMOS camera chip, the use of a multiband emission filter and image splitting optics would be possible and permit simultaneous imaging with multiple wavelengths at no speed penalty. Another possibility would be to alternate the different excitation lasers and use a multiband notch filter to reject the excitation light. The use of a binary ferroelectric SLM in this implementation also is not optimal. The diffraction efficiency of such an SLM is very low, so most of the incident light is in the zero order reflection, which is filtered out by the spatial mask. For applications requiring very high frame rates, the imaging speed is therefore limited by the output power of the laser diodes. The SLM also introduces some ellipticity in the polarization for wavelengths away from the 550 nm design wavelength where the pixels do not operate as ideal half wave plates. Although this could be compensated for by using an additional LCVR, the ideal solution may be the use of a digital micro-mirror device (DMD) as a pattern generator.
Possible modifications
The setup presented here is flexible and more easily modified than commercial instruments so other imaging modalities such as 3D-SIM, fast 2D-SIM, multifocal SIM (MSIM) and non-linear SIM (NL-SIM) can be implemented21,34,35.
2D-SIM can be well suited for imaging relatively flat, fast moving structures such as the peripheral endoplasmic reticulum. The peripheral ER lies deeper within the cell than can be illuminated using a TIRF evanescent field but due to its flat structure can be imaged using standard 2D-SIM with negligible out-of-focus background. Additionally, the use of improved optical sectioning reconstruction algorithms to suppress out-of-focus light extend the use of 2D-SIM to optically thick samples, albeit where axial resolution doubling is not required21.
In MSIM, the sample is illuminated by a sparse lattice of excitation foci36. This modality can be implemented by simply removing the spatial mask (SM) and replacing it by a polarizer. The SLM now operates as an amplitude modulator. The binary SIM gratings displayed on the SLM can be replaced by a 2D lattice of spots, with the size of the spots chosen to be equal to the size of a diffraction limited focus in the image plane. In Figure 7A, a lattice of 4 x 4 pixel squares is displayed on the SLM (inset) which when demagnified onto the sample generates diffraction limited foci of 150 x 150 nm, given the physical SLM pixel size of 13.62 &#181;m. The excitation foci can then be translated by shifting the lattice pattern on the SLM and this is repeated multiple times in order to illuminate the entire field of view. Images are acquired for each translated pattern position and the stack is post-processed to yield a reconstructed image with improved resolution of up to a factor of <img alt="Equation" src="/files/ftp_upload/53988/53988eq19.jpg" /> and reduced out-of-focus light compared to the equivalent widefield image30. This modality can be useful for imaging thick, dense samples for which standard SIM is unsuited, for example low contrast structures such as stained red blood cells (Figure 7C), although the acquisition time is increased due to the large number of raw frames required per field of view (in this case N = 168).
Finally, the setup can be modified to enable either high-NA linear TIRF-SIM or patterned activation non-linear SIM (PA NL-SIM), as presented recently by Li et al., by use of an ultrahigh 1.7 NA objective or addition of a 405 nm photoactivation laser and careful optimization of the SLM grating patterns35.
Future Applications
SIM is still a rapidly evolving technique and many applications in the life sciences will be enabled in the future. The speed, resolution, and contrast enhancements of the technique and the capability of using standard fluorophores mean that for bioimaging, SIM is set to replace conventional many microscope systems, such as confocal and wide field platforms. Commercial SIM systems are already available today with outstanding technical specifications, however, they are beyond the financial reach of many research laboratories, and, crucially, they are inflexible to be modified and developed to implement the latest research developments in the field. They also lack the essential capability to &#39;be adapted for the experiment at hand&#39;, often a critical bottleneck in cutting edge life science research. The system described here will be particularly well suited to study dynamic processes near the cell surface, for in vitro studies of reconstituted bilayer systems, to study surface chemistry in the materials and physical sciences, e.g. of 2D materials, and many other applications.

Over the last half a decade, super-resolution microscopy has matured and moved from specialist optics labs into the hands of the biologist. Commercial microscope solutions exist for the three main variants for achieving optical super-resolution: single molecule localization microscopy (SMLM), stimulated emission depletion microscopy (STED), and structured illumination microscopy (SIM)1,2. SMLM such as photoactivated localization microscopy (PALM) and stochastic optical reconstruction microscopy (STORM) have been the most popular techniques, largely due to the simplicity of the optical setup and the promise of high spatial resolution, readily down to 20 nm. However super-resolution microscopy via single molecule localization comes with an intrinsic trade-off: the spatial resolution attainable is dependent on accumulating a sufficient number of individual fluorophore localizations, hence limiting the temporal resolution. Imaging dynamic processes in live cells therefore becomes problematic as one must adequately sample the movement of the structure of interest to prevent motion artifacts while also acquiring enough localization events in that time to reconstruct an image. In order to meet these requirements, live cell SMLM demonstrations have obtained the required increase in fluorophore photoswitching rates by greatly increasing the excitation power, and this leads in turn to phototoxicity and oxidative stress, thereby limiting sample survival times and biological relevance3.
A clear advantage of STED over both SIM and SMLM is that it can image with super-resolution in thick samples, for example lateral resolution of around 60 nm was achieved in organotypic brain slices at depths up to 120 &#181;m4. Imaging at such depths with single objective implementations of SMLM or SIM is unfeasible, but becomes possible with either single-molecule light sheet or lattice light sheet microscopy5. Video-rate STED has also been demonstrated and used to map synaptic vesicle mobility, although so far this has been limited to imaging small fields of view6.
For applications in cell biology and molecular self-assembly reactions7-12 that require imaging with high temporal resolution over many time points, structured illumination microscopy (SIM) can be well-suited as it is not dependent on the photophysical properties of a particular fluorescent probe. Despite this inherent advantage of SIM, up to now its use has been mainly confined to imaging fixed cells or slow moving processes. This is due to the limitations of commercially available SIM systems: the acquisition frame rate of these instruments was limited by the rotation speed of the gratings used to generate the required sinusoidal illumination patterns as well as the polarization maintaining optics. The newest generation of commercial SIM instruments are capable of fast imaging but they are prohibitively expensive to all but central imaging facilities.
This protocol presents a guide to the construction of a flexible SIM system for imaging fast processes in thin samples and near the basal surface of living cells. It employs total internal reflection fluorescence (TIRF) to generate an illumination pattern which penetrates no deeper than approximately 150 nm into the sample13 which vastly reduces the out of focus background signal. The idea of combining SIM with TIRF is almost as old as SIM itself14 but was not realized experimentally before 200615. The first in vivo images obtained with TIRF-SIM were reported in 200916 achieving frame rates of 11 Hz to visualize tubulin and kinesin dynamics, and two color TIRF-SIM systems have been presented17,18. Most recently, a guide for the construction and use of a single color two-beam SIM system was presented featuring frame-rates of up to 18 Hz19,20.
The set-up presented here is capable of SIM super-resolution imaging at 20 Hz in three colors, two of which can be operated in TIRF-SIM. The whole system is constructed around an inverted microscope frame and uses a motorized xy translation stage with a piezo-actuated z stage. To generate the sinusoidal excitation patterns required for TIRF-SIM, the system presented uses a ferroelectric spatial light modulator (SLM). Binary grating patterns are displayed on the SLM and the resulting &#177;1 diffraction orders are filtered, relayed and focused into the TIR ring of the objective lens. The necessary phase shifts and rotations of the gratings are applied by changing the displayed SLM image. This protocol describes how to build and align such an excitation path, details the alignment of the emission path, and presents test samples for ensuring optimal alignment. It also describes the issues and challenges particular to high speed TIRF-SIM regarding polarization control and synchronization of components.
Design Considerations and Constraints
Before assembling the TIRF-SIM system presented in this protocol, there are several design constraints to consider which determine the choice of optical components. All abbreviations of optical components refer to Figure 1.
Spatial Light Modulator (SLM)
A binary ferroelectric SLM is used in this setup as it is capable of sub-millisecond pattern switching. Grayscale nematic SLMs may be used but these offer greatly reduced switching times. Each on or off pixel in a binary phase SLM will impart either a &#960; or 0 phase offset to the incident plane wavefront, therefore if a periodic grating pattern is displayed on the SLM it will operate as a phase diffraction grating.
Total Internal Reflection (TIR)
To achieve TIR and produce an evanescent field, the incident angle of the excitation beams at the glass-sample interface must be greater than the critical angle <img alt="Equation" src="/files/ftp_upload/53988/53988eq1.jpg" />. This sets the minimum incident angle required, and hence also the maximum spacing, or period, of the evanescent illumination pattern. The maximum incident angle <img alt="Equation" src="/files/ftp_upload/53988/53988eq2.jpg" /> (the acceptance angle) is limited by the numerical aperture (NA) of the objective lens which can be calculated from the definition <img alt="Equation" src="/files/ftp_upload/53988/53988eq3.jpg" />. This determines the minimum pattern spacing achievable according to the Abbe formula <img alt="Equation" src="/files/ftp_upload/53988/53988eq4.jpg" /> which links NA and wavelength <img alt="Equation" src="/files/ftp_upload/53988/53988eq5.jpg" /> to the minimum pattern spacing <img alt="Equation" src="/files/ftp_upload/53988/53988eq6.jpg" />. In practice, a 1.49 NA oil immersion TIRF objective yields a maximum angle of incidence of around 79&#176; and a minimum pattern period on the sample of 164 nm using an excitation wavelength of 488 nm. These two angles define a ring in the back aperture of the objective over which the instrument achieves TIR illumination (i.e. the TIR ring) and in which the two excitation foci must be accurately positioned and precisely rotated to generate each illumination pattern.
Reconstruction of TIRF-SIM images requires the acquisition of a minimum of three phase shifts per pattern rotation therefore the SLM pattern period must be divisible by 3 (see Figure 1). For example, a period of 9 pixels for 488 nm illumination and 12 pixels for 640 nm illumination. For a comprehensive discussion of SLM pattern design, including sub-pixel optimization of pattern spacing using sheared gratings, see the previous work of Kner et al.16 and Lu-Walther et al.20 The position of the two excitation foci must be inside the TIR ring for all wavelengths, however the diffraction angle of the &#177;1 orders from the SLM is wavelength dependent. For standard SIM, multicolor imaging can be achieved by optimizing the grating period for the longest wavelength, and tolerating a loss in performance for the shorter channels. For TIRF-SIM however, optimizing for one wavelength means that the other wavelength foci are no longer within the TIR ring. For example, using a grating period of 9 pixels is sufficient to provide TIRF for 488 nm, as the foci are at 95% of the diameter of the back aperture and within the TIR ring, but for 640 nm this period would position the foci outside the aperture. For this reason different pixel pattern spacings must be used for each excitation wavelength.
The alignment of the TIRF-SIM excitation path is extremely sensitive to small changes in the position of the dichroic mirror (DM4 in Figure&#160;1) in the microscope body, much more so than in conventional SIM. Use of a rotating filter cube turret is not recommended, instead use a single, multi-band dichroic mirror, which is kept in a fixed position and designed specifically for the excitation wavelengths used. It is essential that only the highest quality dichroic mirrors are used. These require thick substrates of at least 3 mm, and are often designated as &#34;imaging flat&#34; by manufacturers. All other substrates lead to intolerable aberration and image degradation in TIRF-SIM.
Polarization Control
To achieve TIRF-SIM it is essential to rotate the polarization state of the excitation light in synchronicity with the illumination pattern such that it remains azimuthally polarized in the objective pupil plane with respect to the optical axis (i.e. s-polarized). Alignment of the polarization control optics will depend on the specific optical element employed, for example a Pockels cell21, or a half wave plate in a motorized rotation stage22. In this protocol a custom liquid crystal variable retarder (LCVR) is used, designed to provide full-wave (2&#960;) retardance over the wavelength range 488 to 640 nm as it allows fast (~msec) switching. If using a liquid crystal retarder it is essential to use a high quality component: standard components are typically not stable enough to give a constant retardance over the length of the camera exposure time which leads to a blurring out of the illumination pattern and low modulation contrast. Liquid crystal retarders are also strongly temperature dependent and require built in temperature control.
Synchronization
The lasers must be synchronized with the SLM. Binary ferroelectric SLMs are internally balanced by switching between an on state and off state. The pixels only act as half wave plates in either their on or off state, but not during the interframe switching time. Therefore the lasers should only be switched on during on/off states via the LED Enable signal from the SLM to prevent a reduction in pattern contrast due to the intermediate state of the pixels. An acousto-optic modulator (AOM) could alternatively be used as a fast shutter if the lasers cannot be digitally modulated.
Choice of Lenses
Based on these constraints, the required demagnification of the SLM plane onto the sample plane to produce the desired illumination patterns can be determined. This allows calculation of the focal lengths of the two lenses L3 and L4 in the image relay telescope and the excitation condenser lens L5. In this system a 100X/1.49NA oil immersion objective lens is used with 488 nm and 640 nm excitation, hence uses focal lengths of 300 and 140 mm for L4 and L3, and 300 mm for L5, giving a total demagnification of 357X, equivalent to an SLM pixel size of 38 nm at the sample plane. Using this combination of lenses, SLM grating periods of 9 for 488 nm illumination and 12 pixels for 640 nm give pattern spacings of 172 and 229 nm at the sample, corresponding to angles of incidence of 70&#176; and 67&#176; respectively. For a glass-water interface, the critical angle is 61&#176;, and is independent of wavelength, therefore these two pattern spacings allow TIRF excitation for both wavelengths. An objective lens equipped with a correction collar is useful for correction of spherical aberrations introduced by variations in coverslip thickness, or if operating at 37 &#176;C.
Image Reconstruction
Once raw SIM data has been acquired it is a matter of computational effort to generate super-resolved images in a two-step process. Firstly, the illumination pattern has to be determined for every image and secondly, the components of the SIM spectrum must be separated and recombined appropriately as to double the effective OTF support (see Figure 6, insets).
Precise knowledge of the projected illumination patterns is paramount, as the super-resolved frequency components have to be unmixed as accurately as possible to prevent artifacts caused by the residual parts of overlapping components. We determine the illumination pattern parameters a posteriori from the raw image data following the procedure introduced by Gustafsson et al.23 In short, a set of illumination parameters that describes a normalized two-dimensional sinusoid has to be found for each of the <img alt="Equation" src="/files/ftp_upload/53988/53988eq7.jpg" /> excitation patterns <img alt="Equation" src="/files/ftp_upload/53988/53988eq8.jpg" />:
<img alt="Equation" src="/files/ftp_upload/53988/53988eq9.jpg" />
Hereby <img alt="Equation" src="/files/ftp_upload/53988/53988eq10.jpg" /> and <img alt="Equation" src="/files/ftp_upload/53988/53988eq11.jpg" /> describe the fringe contrast and the pattern starting phase of each individual image m respectively. The components of the wave vector, <img alt="Equation" src="/files/ftp_upload/53988/53988eq12.jpg" /> and <img alt="Equation" src="/files/ftp_upload/53988/53988eq13.jpg" /> , only change with different orientations <img alt="Equation" src="/files/ftp_upload/53988/53988eq14.jpg" /> of the pattern and can assumed to be otherwise constant. To coarsely determine the components of the wave vector a cross correlation of raw image spectra is performed, which is refined by applying subpixel shifts to one of the cross-correlated images as to optimize the overlap. This is done via multiplication of real-space phase gradients <img alt="Equation" src="/files/ftp_upload/53988/53988eq15.jpg" /> that induce a subpixel shift in frequency-space. Note that it is useful to have a good estimate of the wave-vectors prior to the actual pattern estimation and this can be found by imaging a fluorescent bead layer.
As the phase step between shifted patterns is <img alt="Equation" src="/files/ftp_upload/53988/53988eq16.jpg" />, i.e. <img alt="Equation" src="/files/ftp_upload/53988/53988eq17.jpg" />, the separation of frequency components can be performed by a Fourier transform along the &#34;phase axis&#34;. The global phase <img alt="Equation" src="/files/ftp_upload/53988/53988eq18.jpg" /> and the fringe contrast <img alt="Equation" src="/files/ftp_upload/53988/53988eq10.jpg" /> can then be determined using complex linear regression of different components. The individual separated components are then combined using a generalised Wiener filter. For a detailed description of both parameter extraction and implementation of the generalized Wiener filter we refer the reader to Gustafsson et al.23 where the same algorithm is used.

<table><tbody><tr><td>488 nm laser</td><td>Toptica</td><td>iBeam SMART</td><td>with digital modulation</td></tr><tr><td>561 nm laser</td><td>Coherent</td><td>OBIS LS</td><td>with digital modulation</td></tr><tr><td>640 nm laser</td><td>Cobolt</td><td>MLD</td><td>with digital modulation</td></tr><tr><td>Long-pass dichroic mirrors&amp;nbsp;</td><td>Thorlabs</td><td></td><td>for combining excitation beams</td></tr><tr><td>Quad band dichroic mirror</td><td>Chroma</td><td>ZT405/488/561/640rpc</td><td>3 mm thick, TIRF imaging flat, mounted in Olympus BX filter cube</td></tr><tr><td>Quad band dichroic mirror</td><td>Chroma</td><td>ZT405/488/561/640rpc</td><td>From same batch as above, 25 x 25 mm</td></tr><tr><td>1&quot; square kinematic mount</td><td>Edmund Optics</td><td>58-860</td><td></td></tr><tr><td>Glan-Taylor calcite polarizers</td><td>Thorlabs</td><td>GT5-A</td><td>For alignment of LCVR</td></tr><tr><td>Glan-Taylor mount</td><td>Thorlabs</td><td>SM05PM5</td><td></td></tr><tr><td>Achromatic half wave plate</td><td>Thorlabs</td><td>AHWP05M-600</td><td>400-800 nm</td></tr><tr><td>Rotation cage mount</td><td>Thorlabs</td><td>CRM1/M</td><td>For HWP</td></tr><tr><td>Liquid Crystal Variable Retarder</td><td>Meadowlark Optics</td><td>SWIFT</td><td>Custom built to provide full wave retardance over the range 488 to 640 nm.</td></tr><tr><td>LCVR controller</td><td>Meadowlark Optics</td><td>D3060HV</td><td>Two channel high voltage controller for liquid crystal retarders</td></tr><tr><td>Achromatic quarter wave plate</td><td>Meadowlark Optics</td><td>AQM-100-0545</td><td></td></tr><tr><td>Rotation cage mount</td><td>Thorlabs</td><td>CRM1P/M</td><td>For QWP</td></tr><tr><td>10 mm achromatic doublet</td><td>Thorlabs</td><td>AC080-010-A-ML</td><td>For beam expander</td></tr><tr><td>200 mm achromatic doublet</td><td>Thorlabs</td><td>AC254-200-A-ML</td><td>For beam expander</td></tr><tr><td>Cage XY Translators</td><td>Thorlabs</td><td>CXY1</td><td></td></tr><tr><td>Ferroelectric spatial light modulator</td><td>Forth Dimension Displays</td><td>M0787-00249&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;</td><td>SXGA-3DM (IFF) Microdisplay Type M249, 1,280 x 1,024 pixels, with driver board</td></tr><tr><td>SLM mounting frame</td><td>Forth Dimension Displays</td><td>M0787-10014</td><td>Fixed to custom built aluminium mount</td></tr><tr><td>&amp;Oslash;50.8 mm Gimbal Mirror Mount</td><td>Thorlabs</td><td>GM200/M</td><td>For SLM mounting</td></tr><tr><td>Two-Axis Linear Translation Stage with Rotating Platform</td><td>Thorlabs</td><td>XYR1/M</td><td>For SLM mounting</td></tr><tr><td>Rail carrier</td><td>Newport</td><td>M-PRC-3</td><td>For SLM mounting</td></tr><tr><td>Precision Optical Rail</td><td>Newport</td><td>PRL-6</td><td>For SLM mounting</td></tr><tr><td>300 mm achromatic doublet lens</td><td>Qioptiq</td><td>G322 273 322&amp;nbsp;</td><td>f = 300 mm, 31.5 mm diameter</td></tr><tr><td>140 mm achromatic doublet lens</td><td>Qioptiq</td><td>G322 239 322</td><td>f = 140 mm, 31.5 mm diameter</td></tr><tr><td>Precision XY Translation Mounts</td><td>Thorlabs</td><td>LM2XY</td><td></td></tr><tr><td>Lens Mounting Adapters</td><td>Thorlabs</td><td>SM2AD32</td><td>For mounting 31.5 mm lenses in 2&quot; mounts</td></tr><tr><td>Translation stages</td><td>Comar</td><td>12XT65</td><td>Dovetail, side drive</td></tr><tr><td>XY Translator with Differential Drives</td><td>Thorlabs</td><td>ST1XY-D/M</td><td>for spatial filter</td></tr><tr><td>Rotation cage mount</td><td>Thorlabs</td><td>CRM1/M</td><td>for spatial filter</td></tr><tr><td>300 mm achromatic doublet</td><td>Thorlabs</td><td>AC508-300-A-ML</td><td>Excitation tube lens</td></tr><tr><td>Automated XY stage with Z-piezo top plate</td><td>ASI</td><td>PZ-2150-XYFT-PZ-IX71&amp;nbsp;</td><td>with MS-2000 controller</td></tr><tr><td>Inverted microscope frame</td><td>Olympus</td><td>IX-71</td><td></td></tr><tr><td>Objective lens</td><td>Olympus</td><td>UAPON100XOTIRF</td><td>100X/1.49NA</td></tr><tr><td>High speed filter wheel</td><td>Prior Scientific</td><td>HF110A</td><td>with Prior ProScan III controller</td></tr><tr><td>Bandpass emission filters</td><td>Semrock</td><td>FF01-525/30, FF01-676/29</td><td></td></tr><tr><td>sCMOS camera</td><td>Hamamatsu</td><td>ORCA Flash v4.0</td><td></td></tr><tr><td>Stage top incubator</td><td>OKO Lab</td><td>H301-K-FRAME</td><td>For live cell imaging, with Bold Line temperature and CO&lt;sub&gt;2&lt;/sub&gt; controllers</td></tr><tr><td>Stainless steel optical posts</td><td>Thorlabs</td><td>TR series</td><td>for mounting optical components</td></tr><tr><td>Post holders</td><td>Thorlabs</td><td>PH series</td><td>for mounting optical components</td></tr><tr><td>Kinematic mirror mounts</td><td>Thorlabs</td><td>KM100</td><td>for mounting 1&quot; mirrors</td></tr><tr><td>Shearing interferometer</td><td>Thorlabs</td><td>SI100</td><td></td></tr><tr><td>100 nm fluorescent microspheres</td><td>Life Technologies</td><td>T-7279</td><td>Tetraspeck</td></tr><tr><td>Rhodamine 6G</td><td>Sigma Aldrich</td><td>83697-250MG</td><td></td></tr><tr><td>8 well glass bottom dishes</td><td>ibidi</td><td>80827</td><td>with #1.5 coverglass</td></tr><tr><td>Nunc Lab-Tek II Chambered Coverglass</td><td>Thermo Fisher Scientific</td><td>155409</td><td>with #1.5 coverglass</td></tr><tr><td>0.01 mm microscope reticle slide</td><td>EMS</td><td>68039-22</td><td></td></tr><tr><td>CellLight Tubulin-GFP, BacMam 2.0</td><td>Thermo Fisher Scientific</td><td>C10613</td><td></td></tr></tbody></table>

a guide to structured illumination tirf microscopy at high speed with multiple colors

Optical super-resolution imaging with structured illumination microscopy (SIM) is a key technology for the visualization of processes at the molecular level in the chemical and biomedical sciences. Although commercial SIM systems are available, systems that are custom designed in the laboratory can outperform commercial systems, the latter typically designed for ease of use and general purpose applications, both in terms of imaging fidelity and speed. This article presents an in-depth guide to building a SIM system that uses total internal reflection (TIR) illumination and is capable of imaging at up to 10 Hz in three colors at a resolution reaching 100 nm. Due to the combination of SIM and TIRF, the system provides better image contrast than rival technologies. To achieve these specifications, several optical elements are used to enable automated control over the polarization state and spatial structure of the illumination light for all available excitation wavelengths. Full details on hardware implementation and control are given to achieve synchronization between excitation light pattern generation, wavelength, polarization state, and camera control with an emphasis on achieving maximum acquisition frame rate. A step-by-step protocol for system alignment and calibration is presented and the achievable resolution improvement is validated on ideal test samples. The capability for video-rate super-resolution imaging is demonstrated with living cells.

Watch this Scientific Journal Video about A Guide to Structured Illumination TIRF Microscopy at High Speed with Multiple Colors at JoVE.com

A Guide to Structured Illumination TIRF Microscopy at High Speed with Multiple Colors

This article provides an in depth guide for the assembly and operation of a structured illumination microscope operating with total internal reflection fluorescence illumination (TIRF-SIM) to image dynamic biological processes with optical super-resolution in multiple colors.

Optical super-resolution imaging with structured illumination microscopy (SIM) is a key technology for the visualization of processes at the molecular ...

a-guide-to-structured-illumination-tirf-microscopy-at-high-speed-with

Universidad San Sebastian

Research

JoVE Journal

Bioengineering

25.1K Views.  University of Cambridge. This article provides an in depth guide for the assembly and operation of a structured illumination microscope operating with total internal reflection fluorescence illumination (TIRF-SIM) to image dynamic biological processes with optical super-resolution in multiple colors.

Video: A Guide to Structured Illumination TIRF Microscopy at High Speed with Multiple Colors

Visualization of Cortex Organization and Dynamics in Microorganisms, using Total Internal Reflection Fluorescence Microscopy

TIRF microscopy has emerged as a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules in vitro and in living cells1. The optical phenomenon of total internal reflection occurs when light passes from a medium with high refractive index into a medium with low refractive index at an angle larger than a characteristic critical angle (i.e. closer to being parallel with the boundary). Although all light is reflected back under such conditions, an evanescent wave is created that propagates across and along the boundary, which decays exponentially with distance, and only penetrates sample areas that are 100-200 nm near the interface2. In addition to providing superior axial resolution, the reduced excitation of out of focus fluorophores creates a very high signal to noise ratios and minimizes damaging effects of photobleaching2,3. Being a widefield technique, TIRF also allows faster image acquisition than most scanning based confocal setups.
At first glance, the low penetration depth of TIRF seems to be incompatible with imaging of bacterial and fungal cells, which are often surrounded by thick cell walls. On the contrary, we have found that the cell walls of yeast and bacterial cells actually improve the usability of TIRF and increase the range of observable structures4-8. Many cellular processes can therefore be directly accessed by TIRF in small, single-cell microorganisms, which often offer powerful genetic manipulation techniques. This allows us to perform in vivo biochemistry experiments, where kinetics of protein interactions and activities can be directly assessed in living cells.
We describe here the individual steps required to obtain high quality TIRF images for Saccharomyces cerevisiae or Bacillus subtilis cells. We point out various problems that can affect TIRF visualization of fluorescent probes in cells and illustrate the procedure with several application examples. Finally, we demonstrate how TIRF images can be further improved using established image restoration techniques.

Total Internal Reflection Fluorescence (TIRF) microscopy is a powerful approach to observe structures close to the cell surface at high contrast and temporal resolution. We demonstrate how TIRF can be employed to study protein dynamics at the cortex of cell wall-enclosed bacterial and fungal cells.

TIRF microscopy has emerged as  a powerful imaging technology to study spatio-temporal dynamics of fluorescent molecules  in vitro and in living cells1. ...

A TIRF Microscopy Technique for Real-time, Simultaneous Imaging of the TCR and its Associated Signaling Proteins

Signaling is initiated through the T Cell Receptor (TCR) when it is engaged by antigenic peptide fragments bound by Major Histocompatibility Complex (pMHC) proteins expressed on the surface of antigen presenting cells (APCs). The TCR complex is composed of the ligand binding TCR&alpha;&beta; heterodimer that associates non-covalently with CD3 dimers (the &#949;&delta; and &#949;&gamma; heterodimers and the &zeta;&zeta; homodimer)1. Upon engagement of the receptor, the CD3 &zeta; chains are phosphorylated by the Src family kinase, Lck. This leads to the recruitment of the Syk family kinase, Zap70, which is then phosphorylated and activated by Lck. After that, Zap70 phosphorylates the adapter proteins LAT and SLP76, initiating the formation of the proximal signaling complex containing a large number of different signaling molecules2. 
The formation of this complex eventually results in calcium and Ras-dependent transcription factor activation and the consequent initiation of a complex series of gene expression programs that give rise to T cell differentiation2. TCR signals (and the resulting state of differentiation) are modulated by many other factors, including antigen potency and crosstalk with co-stimulatory/co-inhibitory, chemokine, and cytokine receptors 3-4. Studying the spatial and temporal organization of the proximal signaling complex under various stimulation conditions is, therefore, key to understanding the TCR signaling pathway as well as its regulation by other signaling pathways.
One very useful model system to study signaling initiated by the TCR at the plasma membrane in T cells is glass-supported lipid bilayers, as described previously5-6. They can be utilized to present antigenic pMHC complexes, adhesion, and co-stimulatory molecules to T cells-serving as artificial APCs. By imaging the T cells interacting with the lipid bilayer using total internal reflection fluorescence microscopy (TIRFM), we can restrict the excitation to within 100 nm of the space between the glass and the cell surface 7-8. This allows us to image primarily the signaling events occurring at the plasma membrane. As we are interested in imaging the recruitment of signaling proteins to the TCR complex, we describe a two-camera TIRF imaging system wherein the TCR, labeled with fluorescent Fab (fragment antigen binding) fragments of the H57 antibody (purified from hybridoma H57-597, ATCC, ATCC Number:HB-218) which is specific for TCR&beta;, and signaling proteins, tagged with GFP, may be imaged simultaneously and in real time. This strategy is necessary due to the highly dynamic nature of both the T cells and of the signaling events that are occurring at the TCR. This imaging modality has allowed researchers to image single ligands 9-11 as well as recruitment of signaling molecules to activated receptors and is an excellent system to study biochemistry in-situ12-16.

The compartmentalization of proteins either within the plasma membrane or into intracellular locations is one regulatory mechanism that can greatly influence signaling outcomes; hence, to understand signaling it is important to study the spatial and temporal behavior of the proteins involved. We describe here a TIRF microscopy based system to study signal transduction in T cells, but is broadly applicable.

Signaling is initiated through the T Cell Receptor (TCR) when it is engaged by antigenic peptide fragments bound by Major Histocompatibility Complex ...

Immunology and Infection

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 ...

Biochemistry

Cortical Actin Flow in T Cells Quantified by Spatio-temporal Image Correlation Spectroscopy of Structured Illumination Microscopy Data

Filamentous-actin plays a crucial role in a majority of cell processes including motility and, in immune cells, the formation of a key cell-cell interaction known as the immunological synapse. F-actin is also speculated to play a role in regulating molecular distributions at the membrane of cells including sub-membranous vesicle dynamics and protein clustering. While standard light microscope techniques allow generalized and diffraction-limited observations to be made, many cellular and molecular events including clustering and molecular flow occur in populations at length-scales far below the resolving power of standard light microscopy. By combining total internal reflection fluorescence with the super resolution imaging method structured illumination microscopy, the two-dimensional molecular flow of F-actin at the immune synapse of T cells was recorded. Spatio-temporal image correlation spectroscopy (STICS) was then applied, which generates quantifiable results in the form of velocity histograms and vector maps representing flow directionality and magnitude. This protocol describes the combination of super-resolution imaging and STICS techniques to generate flow vectors at sub-diffraction levels of detail. This technique was used to confirm an actin flow that is symmetrically retrograde and centripetal throughout the periphery of T cells upon synapse formation.

To investigate flow velocities and directionality of filamentous-actin at the T cell immunological synapse, live-cell super-resolution imaging is combined with total internal reflection fluorescence and quantified with spatio-temporal image correlation spectroscopy.

Filamentous-actin plays a crucial role in a majority of cell processes including motility and, in immune cells, the formation of a key cell-cell ...

Simultaneous Interference Reflection and Total Internal Reflection Fluorescence Microscopy for Imaging Dynamic Microtubules and Associated Proteins

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. Total-internal-reflection-fluorescence (TIRF) microscopy has a high signal-to-background ratio, but it suffers from photobleaching and photodamage of the fluorescent proteins. Label-free techniques such as interference reflection microscopy (IRM) and interferometric scattering microscopy (iSCAT) circumvent the problem of photobleaching but cannot readily visualize single molecules. This paper presents a protocol for combining IRM with a commercial TIRF microscope for the simultaneous imaging of microtubule-associated proteins (MAPs) and dynamic microtubules in vitro. This protocol allows for high-speed observation of MAPs interacting with dynamic microtubules. This improves on existing two-color TIRF setups by eliminating both the need for microtubule labeling and the need for several additional optical components, such as a second excitation laser. Both channels are imaged on the same camera chip to avoid image registration and frame synchronization problems. This setup is demonstrated by visualizing single kinesin molecules walking on dynamic microtubules.

We present a protocol for implementing interference-reflection microscopy and total-internal-reflection-fluorescence microscopy for the simultaneous imaging of dynamic microtubules and fluorescently labeled microtubule-associated proteins.

Several techniques have been employed for the direct visualization of cytoskeletal filaments and their associated proteins. ...

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 ...

Visualizing Adhesion Formation in Cells by Means of Advanced Spinning Disk-Total Internal Reflection Fluorescence Microscopy

In living cells, processes such as adhesion formation involve extensive structural changes in the plasma membrane and the cell interior. In order to visualize these highly dynamic events, two complementary light microscopy techniques that allow fast imaging of live samples were combined: spinning disk microscopy (SD) for fast and high-resolution volume recording and total internal reflection fluorescence (TIRF) microscopy for precise localization and visualization of the plasma membrane. A comprehensive and complete imaging protocol will be shown for guiding through sample preparation, microscope calibration, image formation and acquisition, resulting in multi-color SD-TIRF live imaging series with high spatio-temporal resolution. All necessary image post-processing steps to generate multi-dimensional live imaging datasets, i.e. registration and combination of the individual channels, are provided in a self-written macro for the open source software ImageJ. The imaging of fluorescent proteins during initiation and maturation of adhesion complexes, as well as the formation of the actin cytoskeletal network, was used as a proof of principle for this novel approach. The combination of high resolution 3D microscopy and TIRF provided a detailed description of these complex processes within the cellular environment and, at the same time, precise localization of the membrane-associated molecules detected with a high signal-to-background ratio.

An advanced microscope that permit fast and high-resolution imaging of both, the isolated plasma membrane and the surrounding intracellular volume, will be presented. The integration of spinning disk and total internal reflection fluorescence microscopy in one setup allows live imaging experiments at high acquisition rates up to 3.5 s per image stack.

In living cells, processes such as adhesion formation involve extensive structural changes in the plasma membrane and the cell interior. In order to ...

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

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

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

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

Biology

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, ...

Multi-color Localization Microscopy of Single Membrane Proteins in Organelles of Live Mammalian Cells

Knowledge about the localization of proteins in cellular subcompartments is crucial to understand their specific function. Here, we present a super-resolution technique that allows for the determination of the microcompartments that are accessible for proteins by generating localization and tracking maps of these proteins. Moreover, by multi-color localization microscopy, the localization and tracking profiles of proteins in different subcompartments are obtained simultaneously. The technique is specific for live cells and is based on the repetitive imaging of single mobile membrane proteins. Proteins of interest are genetically fused with specific, so-called self-labeling tags. These tags are enzymes that react with a substrate in a covalent manner. Conjugated to these substrates are fluorescent dyes. Reaction of the enzyme-tagged proteins with the fluorescence labeled substrates results in labeled proteins. Here, Tetramethylrhodamine (TMR) and Silicon Rhodamine (SiR) are used as fluorescent dyes attached to the substrates of the enzymes. By using substrate concentrations in the pM to nM range, sub-stoichiometric labeling is achieved that results in distinct signals. These signals are localized with ~15&#8211;27 nm precision. The technique allows for multi-color imaging of single molecules, whereby the number of colors is limited by the available membrane-permeable dyes and the repertoire of self-labeling enzymes. We show the feasibility of the technique by determining the localization of the quality control enzyme (Pten)-induced kinase 1 (PINK1) in different mitochondrial compartments during its processing in relation to other membrane proteins. The test for true physical interactions between differently labeled single proteins by single molecule FRET or co-tracking is restricted, though, because the low labeling degrees decrease the probability for having two adjacent proteins labeled at the same time. While the technique is strong for imaging proteins in membrane compartments, in most cases it is not appropriate to determine the localization of highly mobile soluble proteins.

Here, we present a protocol for multi-color localization of single membrane proteins in organelles of live cells. To attach fluorophores, self-labeling proteins are used. Proteins, located in different membranes compartments of the same organelle, can be localized with a precision of ~18 nm.