Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Summary

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules 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 within single cells.

Abstract

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 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.

Introduction

While cellular structures exist on a wide range of spatial scales, fluorescence imaging of cellular organization on length scales smaller than ~250 nm is restricted in conventional microscopy due to the physical constraint of the diffraction limit. This limit was overcome with the advent of fluorescence photoactivation localization microscopy (FPALM¹) and similar techniques^2,3, which can localize large numbers of individual molecules with precision of ~10 nm, to generate images with resolution of a few tens of nanometers. FPALM is based on using optical control to activate and inactivate subsets of molecules (for a full description of FPALM, and instructions on how to implement this imaging system, see Gould et al.⁴). This technique allows for the spatial distributions of whole populations of single molecules to be mapped, thereby elucidating biological structures across length scales spanning from tens of nanometers to tens of microns. Localization-based super-resolution microscopy (hereto referred to as localization microscopy) has now been adapted to address a range of biological questions, with technological developments permitting, for example, the imaging of individual molecular orientations with polarization FPALM, or P-FPALM⁵, the fluorescence imaging of single molecules in three dimensions with Biplane FPALM⁶ or other techniques^7-9, and the super-resolution fluorescence imaging of single molecules in living cells^10-12. Localization microscopy has also been applied to the imaging of multiple species in fixed cells^13-16. Recently, three protein species have been simultaneously imaged with FPALM in both fixed and living cells¹⁷. Localization microscopy can image samples labeled in a variety of ways: examples include proteins expressed with PAFP or PSFP fusion tags, antibodies or molecules labeled with caged organic dyes, or conventional organic dyes. While the use of conventional fluorescent dyes allows for the labeling of proteins in the absence of a fusion-protein tag, the conditions generally required for the use of noncaged organic dyes in super-resolution imaging require samples to be immersed in reducing buffers². Additionally, the intracellular delivery of antibody-dye conjugates typically requires cells to be fixed and their membranes permeabilized, or requires that living cells are made permeable through electroporation or some other means. The requirements for reducing buffer conditions and membrane permeabilization limit the suitability of organic dyes for live cell imaging, although recent developments have allowed for effective use of HaloTags and FPALM to image membrane structures¹⁸.

FPALM was the first localization microscopy technique to be applied to live cells¹⁰. In live cells, in addition to providing a time dependent spatial map of the locations of labeled molecules, FPALM can track single molecules over multiple frames, and molecular trajectories determined over timescales of milliseconds¹⁹. Thus, FPALM provides access to fairly short timescales and nanoscale resolution.

Multicolor FPALM can be used for a variety of different probes, including photoactivatable proteins and organic caged or noncaged dyes. We here provide detail on the protocol and setup for the simultaneous imaging of two fluorescent protein species, Dendra2 and PAmCherry. We report the outcomes of imaging PAmCherry conjugated to beta actin (PAmCherry-actin) and Dendra2 conjugated to influenza hemagglutinin (Dendra2-HA) in NIH-3T3 fibroblasts. Components described in the setup can be interchanged for other hardware more suited to the imaging of other probes. Where this is the case, we have tried to be explicit in the text.

Multicolor FPALM is ideal for reporting the spatial distributions of multiple protein species in living or fixed cells. This technique is especially suited to investigating spatial and/or dynamic relationships on nanometer length spatial scales, although images will report localization on a range of length scales, from tens of nanometers up to tens of microns. One major advantage of multicolor FPALM is that the setup is relatively inexpensive to construct, and very flexible for use with various probe combinations. The process of construction and calibration of the system from components also provides considerable understanding of factors which can compromise the quality and interpretability of the data, and so the research outcome. We here detail the methods for the optical setup, sample preparation, and data acquisition of multiple protein species, with PSFP and PAFP fusion constructs, using FPALM. While this protocol describes the analysis of fixed cells, these procedures are readily applicable to the imaging of living cells.

The optical setup here described is ideal for the simultaneous imaging of the PSFP Dendra2 and the PAFP PAmCherry. Many other probes may be used for multicolor imaging; however, the precise components required may vary, depending on the excitation and emission spectra of the chosen probes. Choices of dichroic mirrors, filters, and laser wavelengths should be made based on these considerations.

Protocol

Please note: A diagrammatic representation of optical components referenced in this protocol can be found in Figure 1.

1. Cell Sample Preparation

1. Plate cells at an optimized density (for NIH-3T3 cells, this is roughly 2-5 x 10⁴ cells/cm²) in wells of an 8-well chamber. Cells should be plated in complete media appropriate to the cell type, although media should be made without antibiotics and without phenol red, which contributes to background fluorescence. Note that conditions for cell experimentation, such as the optimal range of passage numbers, may differ for individual cell lines.
2. Incubate cells for 24 hr at 37 °C and 5% CO₂ (or at conditions appropriate for the cell type) to allow cells to adhere to the coverslip. Transfect cells with endotoxin free DNA for each of two protein species constructs (in this case, DNA for PAmCherry-actin and Dendra2-HA). Cover the sample with light impermeable material such as aluminum foil. Transfection should include wells with both DNA constructs, and wells with only one of each of these constructs.
3. Incubate for 4-6 hr, 37 °C and 5% CO₂ (or at conditions appropriate for the cell type), before changing into complete media (with antibiotics, without phenol red) and further incubate for 16-48 hr to allow cells to express the desired proteins.
   1. Cells can be fixed by washing three times with phosphate buffered saline (PBS) then incubate with 4% paraformaldehyde (PFA) (CAUTION: Toxic) in PBS for 15 min at RT, and then washing a further 3x with PBS. However, depending on the proteins of interest, this fixation may result in a sizable pool of tagged molecules which are still motile. To further reduce mobility, alternative fixation methods include the use of chilled 100% methanol, or 0.2% gluteraldehyde and 4% PFA in PBS for >30 min at 25 °C²⁰. In either method, cells should be washed with PBS as above. Note that the use of gluteraldehyde may increase background fluorescence or autofluorescence under some imaging conditions, and may necessitate a post-fixation treatment with sodium borohydride²¹.
4. Sample may be kept at 4 °C, immersed in PBS, sealed in a self-sealing film, for up to 7 days before imaging.

2. Microscope Alignment

1. Place a calibration scale (reticle) onto the microscope stage. Using a 10X objective and the lamp for transmitted light, center the reticle in the center of the field of view (FOV).
2. Köhler Illumination. Adjust the microscope for Köhler illumination²². To begin, close the field aperture and looking through the oculars focus on the reticle. If the edges of the field aperture are out of focus, adjust the height of the condenser until both the field aperture and reticle are in focus.
3. Adjust the lateral position of the field aperture until it is centered with respect to the FOV. Close the field aperture until only the center grid on the reticle is illuminated.
4. For a coarse alignment of camera position (i.e. the first time the setup is aligned), use a high lamp intensity with the camera shutter CLOSED, and do not place any components in box B (Figure 1), into the optical path until step 2.5 is reached. Do not place L2 and L3 into the detection path when first aligning the camera. Roughly center the reticle image on the camera shutter by adjusting the vertical and horizontal position of the camera (Figure 2B). Disable the EM gain, turn off room lights, and open the camera shutter.
5. After reducing the lamp intensity to a level that will not damage the camera sensor, project the light from the reticle image directly onto the camera sensor (Figure 2A). Focus the reticle by adjusting the microscope focus knob while viewing the image in live video mode within the acquisition software. Center the reticle image onto the camera sensor by adjusting the vertical and horizontal position of the camera (Figure 2B).
6. Place L2 and L3 into the detection path between the aperture and the camera (Figure 2C). Align L2 and L3, such that L2 is one focal length from the focal point of the microscope exit port and L3 is one focal length from the camera sensor. The distance between L2 and L3 should ideally be equal to the sum of the focal lengths of L2 and L3, but can be adjusted somewhat to accommodate space constraints. The camera and lenses should be at the same height as the exit port.
7. Note that the light emitted from the microscope should be centered on L2 and L3. Adjust the distance between L2 and the microscope to ensure the reticle image is in sharp focus on both the camera and through the oculars.
8. If necessary, small translations (e.g. <1 mm) of L2 and L3 can be used to center the reticle image onto the camera sensor.
9. Two color module. Once the camera position is optimized, affix components shown in box B (Figure 1) into the detection path. These components can be affixed to a removable mount, so that the entire module can be inserted for multicolor FPALM, or removed for other FPALM applications not requiring it.
10. The first time these components are assembled, adjust the path lengths of each channel to be equal. Project the reticle onto the camera chip, adjust M7 and M9, and/or close the detection aperture (AP) to prevent spatial overlap between the two channels. Focus the image of the reticle in the reflected light channel.
11. If the image in the transmitted light channel is not in focus, translate M9 (and rotate if necessary) until the reticle image is in focus simultaneously in both channels. Note that the two channels should be displaced laterally from one another (Figure 2D). This displacement, whether horizontal or vertical, can affect acquisition speed. For further information, consult the camera user's manual.
12. Record a snapshot of the reticle scale (to later use in calculating the overall magnification). Using the camera software, select the desired region of interest. Higher frame rates will generally be possible for a smaller region of interest.

3. Laser Alignment

1. Turn on the readout and activation lasers. (CAUTION: Lasers should only be used after operators have undergone laser safety training.) All doors to the lab should remain closed, with only trained essential personnel inside the lab whilst lasers are being aligned. Use shutters SH1 and SH2 to block the readout and activation beams respectively when not in use, and ND filters to attenuate laser powers to safe levels (<1 mW). It is helpful to minimize all room lighting during alignment, except for the lighting needed for safety.
2. Block the activation and readout beams. Remove L1 from the laser path.
3. Place a white card flush against M4. Most commercial compound microscopes have a built-in shutter to block incoming illumination. If available, open the microscope shutter, and focus until the reticle image projects onto M4. If an internal microscope shutter is unavailable, use an external shutter in a convenient location that blocks all laser beams from entering the microscope.
4. Centering readout laser in the FOV. Unblock the readout beam. Center the readout beam onto the crosshairs of the reticle image on M4 by adjusting M1.
5. Project the reticle image onto M5, and adjust mirror M4 until the beam is centered on the image crosshairs on M5, such that the readout beam is centered with the reticle crosshair at both M4 and M5. Block the readout beam.
6. Centering activation laser in the FOV. Project the reticle image onto M3, and remove the beam expander (BE) from the laser path. Unblock the activation beam.
7. Adjust M2 to center the activation beam onto the crosshairs of the reticle image on M3. Once centered, replace the BE between M2 and M3, and adjust the position of the BE until the beam is centered on the crosshairs of the reticle image on M3.
8. Using a 10X objective, project the reticle image onto M5, adjusting the microscope focus knob if needed to obtain focus. Adjust the angle of DM1 until the activation beam is centered on the reticle image there. Block both beams.
9. With no objective in place, and the microscope shutter open, project the readout laser through the back aperture of the microscope. (CAUTION: This step creates a laser safety hazard by directing a parallel laser beam in an unblocked vertical path.) Adjust M5 until the beam emerges straight out of the microscope and lands on the ceiling directly above, or is centered on a card placed on the objective mount in the turret.
10. OPTIONAL: Determination of the correct alignment can be facilitated by the use of a sample of dye in solution (in this case, Rhodamine B at ~100 µM in water or methanol with depth of >0.5 cm for visual purposes), placed on the sample stage with the 60X objective lens in place. If the beam is correctly aligned, the objective will project a cone of fluorescence aligned with the axis of the objective and microscope. Small deviations in laser beam placement in the objective back aperture will cause the cone to tip away from a purely vertical alignment.
11. Block both beams. Mount L1 in the laser path at the appropriate distance (i.e. one focal length) from the back aperture of the objective lens. With the 60X objective in place, allow the readout beam to project onto the ceiling. Adjust the horizontal and vertical position of L1 (perpendicular to the direction of laser propagation) until the beam is centered above the microscope. Note: in this step, the beam will form a larger spot than in the previous step.
12. The axial position of L1 and its focal length will affect the size of the illuminated area at the sample. Strictly speaking, calculation of the illumination profile at the sample must take into account diffraction²³. Roughly speaking, however, placing L1 at an axial distance other than one focal length from the objective back focal plane will in many cases cause a smaller, more intense laser illumination profile than when L1 is at exactly one focal length from the back focal plane. A smaller illuminated area can be used to produce a higher laser intensity for certain applications, such as high speed imaging, for example.
13. Measurement of Readout Beam Profile. With L1 in place, place a sample of appropriate concentrated dye solution (in this case, Rhodamine B at ~100 µM in water) onto the stage.
14. With the activation laser blocked, project the readout laser (adjust ND1 to obtain a power <<1 mW for all exposed beams) through the 60X objective and into the dye, and (with the EM gain disabled) send this image to the camera.
15. Focus the objective into the sample. For this step, a large enough aperture is needed to allow imaging of the full beam profile.
16. Translate the AP laterally so that the center of the beam profile and AP are concentric. Using the camera software, choose the region of interest to allow the smallest camera readout region encapsulating both channels. Record these coordinates. Record a single snapshot (this is the readout beam profile).
17. Images which more correctly reflect the laser profile at the focal plane will be obtained when the dye solution sample is as thin as possible. Such a thin sample can be created by placing a drop of ~5 ul of dye solution between a microscope slide and a coverslip.
18. Measurement of Activation Beam Profile. Block the readout laser. Project the activation laser to the sample; and project this image through to the camera.
19. If necessary, use EM gain <100 and adjust DM1 until the beam is centered in each FOV. Record a snapshot of the activation beam profile.
20. Measure the Power of Each Beam. Remove the dye solution and place a power meter sensor over the 60X objective (with no immersion media). Measure the power of each beam (activation and readout) separately. Note that the position of the power meter must be adjusted carefully to ensure that all emitted laser power strikes the power meter sensor.
21. For each laser, use neutral density filters (ND1 and ND2) to adjust power to yield intensities at the sample appropriate for the experiment.
22. Intensity of Readout Laser for Image Acquisition. The readout laser intensity should be made high enough to excite and photobleach single molecules within the time span of a few frames. Typical values are 10³-10⁴ W/cm² (see also Gould et al.⁴). The intensity at the sample is dependent on the size of the image area, so the power required to achieve desired intensity will vary from system to system.
23. Intensity of Activation Laser. The activation laser intensity should be chosen such that the number of active molecules is small (e.g. 1-100) in any given acquisition frame. Roughly speaking, the desired density is reached when the closest distance between active molecules is slightly larger than the diffraction limited resolution (See also Section 6, Imaging). As the population of inactive single molecules decreases, higher intensities of the activation laser are required. Typical intensities are 10^-1-10² W/cm².
24. Optimize the quarter wave plate. The quarter wave plate (QWP) is optional, but increasing the degree of circular polarization of the readout and activation lasers with a QWP can increase molecule density in final images. To optimize the QWP, place a polarizer between the QWP and M5. Block the activation laser. Project the readout laser through to a power meter over the dry 60X objective.
25. Record the angle of the QWP. Adjust the polarizer until the maximum, and then the minimum powers are achieved. Record each of these values and calculate the ratio of minimum/maximum. Adjust the angle of the QWP and repeat these measurements. While it is desirable to obtain values as close to 1.0, a ratio of >0.8 is sufficient for imaging.

4. Creating a Durable Sample of Beads for Channel Alignment

1. Dilute a sample of fluorescent beads (from 40-100 nm in diameter) 1:70 into HPLC grade water. Further dilute this stock solution 1:15 in HPLC water, for a final volume of 200 µl bead suspension in water.
2. Coat a coverslip with liquid poly-L-lysine. Incubate at RT for 30 min. Aspirate to remove the solution, and wash the coverslip three times with HPLC grade water. Aspirate all traces of water from the coverslip and leave to dry at RT.
3. Pipette 200 µl of the bead suspension onto the coverslip. Leave this coverslip for 20 min at RT before washing three times with HPLC water. Alternatively, leave the coverslip O/N at RT to allow the suspension to dry.
4. Using ~20 µl of HPLC water or mounting medium, mount the coverslip onto a glass slide. Seal the periphery of the coverslip with clear nail polish. Once the polish has dried, place the coverslip (and appropriate objective immersion media; either water or oil) onto the 60X objective.

5. Image Acquisition: Imaging Bead Sample for Alignment of Detection Channels

1. Illuminate the bead sample with the readout laser beam at an intensity roughly 10X lower than will be used for imaging. With the EM gain set to 100, project the image to the camera, and adjust the focus until beads are visible in both channels.
2. If beads are dim, either increase the laser power or the EM gain. Minimization of noise in bead images (by detecting a large number of photons, i.e. at least 5,000 in total from each bead) is critical for accurate channel registration. Configure the camera to record 100 frames, with the same exposure time as will be used for subsequent FPALM imaging.
3. Search for regions where beads are distributed in both the center and periphery of the channels, and where bead density is low enough that individual beads are well separated and can be individually identified.
4. Acquire between 10-20 sets of images of different regions at these bead densities. See the results section for details on using the bead images for channel calibration.

6. Image Acquisition: Multicolor FPALM

1. It is important to image cells which have been transfected with only one of each of the constructs, as well as to record images of cells with all constructs. These data will help in establishing the alpha histograms of each of the probes used, and are required for the interpretation of multicolor data.
2. Find cells expressing photoswitchable probes, if in use. Eliminate all room lighting. Project the mercury lamp, via the flip mount (FM), onto the sample (containing transfected cells). Change the turret filter cube to one containing the appropriate dichroic mirror/filter combination to allow for excitation of the prephotoswitched state of the label. For example, for imaging prephotoswitched Dendra2, or probes with a green preswitched emission, one choice for DM4 is a dichroic that reflects blue light (<488 nm) and for F5 is a filter that passes light from ~500-570 nm, while blocking light outside this range.
3. Using the ocular, search for cells which are expressing the prephotoswtiched probe. For example, cells expressing Dendra2 will appear green. Note that not all probes are photoswitchable, and, depending on their preswitched emission spectra, those which are may require combinations of DM4/F5 different from those listed here.
4. Once a cell is chosen, move the FM down to allow the lasers to pass into the microscope (block the mercury light from the path). Change the filter turret to that containing the appropriate dichroic for imaging (in this case, DM2 should reflect both the readout and activation laser, while transmitting longer wavelengths, and F1 should transmit wavelengths to the red of the laser and ideally feature high suppression at the laser wavelength).
5. If cells do not express a photoswitchable probe, search for cells by projecting the image to the camera, and by using the readout laser to illuminate the sample. Single molecules will likely be visible (see Figure 3) in many of the cells.
6. To distinguish transfected cells from background fluorescence (which can still appear as individually flashing molecules), and to confirm that molecules are photoactivatable, briefly illuminate the sample with a low power of the activation laser (typically of order microwatts). The number of photoactivatable molecules visible under the readout beam should dramatically increase and remain high for a short time even after the activation illumination has been once again blocked. Note that different probes may vary considerably in their brightness, photoconversion efficiency, and laser power required for activation^24-27.
7. Prepare the camera software for a kinetic series acquisition by setting EM gain to 200 and choosing the desired number of frames (typically 5,000-10,000), and the exposure time (typically 10-30 msec is appropriate). While the EM gain can be set higher than 200, beyond a certain point increasing the EM gain may increase noise.
8. Block the activation beam. Unblock the readout beam, and project the image of the illuminated cell to the camera.
9. Confirm the cell is transfected (step 6.6). While viewing the cell, adjust focus until the desired focal plane is in view, and molecules are in sharp focus.
10. To choose a focal plane which images near the bottom cellular membrane, shift the focus down until individual molecules are no longer visible. Then, gradually move the focus upward until molecules first become visible.
11. To image near the top cellular membrane, continue to shift the focus up by the desired amount, noting distance using the microscope focus knob or automatic focus. Choosing a focus region between these two limits will result in a region in the middle of the cell being imaged.
12. Unblock the activation beam, and illuminate the sample with a low intensity (use the ND2 filters to attenuate the beam to a very low intensity, roughly <1 W/cm² at the sample).
13. Begin data acquisition. A high density of active molecules is desired; however, it is critical for the analysis steps that these molecules do not overlap spatially.
14. Attempt to maintain a density of visible photoactivatable molecules of ~0.1-1 µm^-2 by adjusting ND2. Typically, for an image region ~10-20 µm in diameter, there will be ~10-100 molecules visible at once (see Figures 3 and 4 for reference). As the number of inactive molecules remaining decreases over the course of the acquisition, the activation laser power may need to gradually increase to maintain the density.
15. If TIRF imaging is desired*, M5 and L1 should both be mounted onto a single translation stage (TS, Figure 1) to be moved laterally (i.e. in a direction perpendicular to the laser just behind the entrance to the microscope). As M5 and L1 are translated, the lasers exiting the objective upward through the sample will gradually tip to one side (CAUTION: laser safety hazard).
16. As the angle of the lasers reaches 90° from the vertical, the emerging laser itself will vanish, the incoming readout laser will be back-reflected, emerging as a beam traveling out of the objective back aperture, anti-parallel to the incoming beam, and displaced to the side.
17. Simultaneously, the background fluorescence will become almost entirely attenuated, and the thickness of the sample region containing discernible, focused single molecules will be greatly reduced.
    *TIRF will permit imaging of a thin section of the sample which is ~100-500 nm above the coverslip. Imaging focal planes which are further into the sample is easily achieved using widefield illumination (Section 3), but is not appropriate for TIRF.
    
18. Upon completion of acquisition, close the microscope shutter immediately, and block both beams. Disable the EM gain, set the camera to record one frame, and set the camera readout region to its maximum size.
19. Block one channel by placing a card over F3 or F4. With a long-pass filter (>580 nm) mounted on the microscope lamp, illuminate the sample and project this image to the camera. Record a snapshot of the cell.
20. OPTIONAL: With one channel still blocked, open the aperture so that a large area of the sample is visible by transmitted light illumination. Record a snapshot. These transmitted light images are very helpful for viewing the context of the corresponding FPALM images.
21. Live cell imaging. To image living cells, transfect cells as per steps 1.1-1.3 but do not fix these samples. Instead, align the setup as described above in full, but before imaging samples, remove them from 37 °C and 5% CO₂, wash 3x in PBS, and immerse sample in imaging media (e.g. PBS with 20 mM glucose). Removal of culture media and washing will reduce the background associated with most cellular media.
22. Samples can be imaged at RT if desired, as fixed cells are, or at 37 °C and 5% CO₂ with the use of an incubation stage mounted onto the microscope stage. A single sample of NIH 3T3 cells should be immersed in imaging media for no more than approximately 1 hr. It may be helpful to monitor how cells respond to immersion in imaging media before preparing for experiments of this kind, to optimize the time of immersion and potentially the composition of the imaging media to reduce perturbation of the cells.

Results

Influenza hemagglutinin (HA) forms clusters on the order of tens of nanometers to micrometers, and these clusters variably colocalize with actin (Figure 5). These spatial distributions corroborate coarser scale imaging of these two proteins²⁸, and the dependence of the HA spatial distributions on actin¹⁹. Multicolor FPALM images can be further used to describe the density, area and perimeter of these clusters, and the degree of colocalization between the two species at both the nano...

Discussion

Localization-based super-resolution imaging provides many powerful capabilities for biological imaging. The route from individual optical components placed on the table to a functional super-resolution microscope capable of simultaneously imaging multiple fluorescent species in a biological sample presents a number of challenges. Some aspects of the alignment are more critical than others; we endeavor below to provide guidance to prospective users dealing with the most difficult aspects of the route.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
LabTek II chambers	Nunc
Fluorescent beads	Invitrogen	F-8801	Beads for calibration
Tetraspeck beads	Invitrogen	T-7279	Four color beads for calibration
Objective immersion oil	Zeiss	518F	Immersion oil for high NA objective (dependent on choice of objective)
HPLC water	Fisher Scientific	W5-4
Media	ATCC	30-2003	Or Cellgro 10-090
Antibiotics	GIBCO	15070-063
serum	Thermo Scientific	SH30087.03
Lipofectamine	Invitrogen	52887
Optimem I	GIBCO	11058-021
Trypsin	MPBiomedicals	1689149
paraformaldehyde	Fisher Scientific	AA433689M	CAUTION: Toxic