Conducting Multiple Imaging Modes with One Fluorescence Microscope

Podsumowanie

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.

Streszczenie

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 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.

Wprowadzenie

Fluorescence microscopes are important tools for the modern biological science research and fluorescent imaging is routinely performed in many biology laboratories. By tagging biomolecules of interest with fluorophores, we can directly visualize them under the microscope and record the time-dependent changes in localization, conformation, interaction, and assembly state in vivo or in vitro. Conventional fluorescence microscopes have a diffraction-limited spatial resolution, which is ~200 - 300 nm in the lateral direction and ~500 - 700 nm in the axial direction^1,2, and are, therefore, limited to imaging at the 100s of nanometers-to-micron scale. In order to reveal finer details in the molecular assembly or organization, various SR microscopies that can break the diffraction limit have been developed. Strategies used to achieve SR include non-linear optical effects, such as stimulated emission depletion (STED) microscopy^3,4 and structured illumination microscopy (SIM)^5,6,7, stochastic detection of single molecules, such as stochastic optical reconstruction microscopy (STORM)⁸ and photoactivated localization microscopy (PALM)⁹, and a combination of both, such as MINFLUX¹⁰. Among these SR microscopies, single-molecule detection-based SR microscopes can be relatively easily modified from a single-molecule microscope set-up. With repetitive activation and imaging of photoactivatable fluorescent proteins (FPs) or photo-switchable dyes tagged on biomolecules of interest, spatial resolution can reach 10 - 20 nm¹¹. To gain information on molecular interactions and conformational dynamics in real-time, angstrom-to-nanometer resolution is required. smFRET^12,13 is one approach to achieve this resolution. Generally, depending on the biological questions of interest, imaging methods with different spatial resolutions are needed.

Typically, for each type of imaging, specific excitation and/or emission optical configuration is needed. For instance, one of the most commonly used illumination methods for single-molecule detection is through total internal reflection (TIR), in which a specific excitation angle needs to be achieved either through a prism or through the objective lens. For smFRET detection, emissions from both donor and acceptor dyes need to be spatially separated and directed to different parts of the electron-multiplying, charge-coupled device (EMCCD), which can be achieved with a set of mirrors and dichroic beam splitters placed in the emission path. For three-dimensional (3-D) SR imaging, an optical component, such as a cylindrical lens¹⁴, is needed to cause an astigmatism effect in the emission path. Therefore, homebuilt or commercially available integrated microscopes are, usually, functionally specialized for each type of imaging method and are not flexible to switch between different imaging methods on the same set-up. Here we present a cost-effective, hybrid system that provides adjustable and reproducible switches between three different imaging methods: conventional epi-fluorescent imaging with diffraction-limited resolution, single-molecule detection-based SR imaging, and multi-color single-molecule detection, including smFRET imaging (Figure 1A). Specifically, the set-up presented here contains fiber-coupled input lasers for multi-color excitation and a commercial illumination arm in the excitation path, which allows programmed control of the excitation angle, to switch between epi-mode and TIR mode. In the emission path, a removable homebuilt cylindrical lens cassette is placed within the microscope body for 3-D SR imaging, and a commercial beam splitter is placed before an EMCCD camera that can be selectively enabled to detect multiple emission channels simultaneously.

Protokół

1. Microscope Design and Assembly

1. Excitation path
   NOTE: The excitation path includes lasers, differential interference contrast (DIC) components, the microscope body, and its illumination arm.
   1. Prepare a vibration-isolated optical table. For example, a structural damping table of 48 x 96 x 12’’ gives enough space for all the components.
      NOTE: Build the set-up in a room with temperature control (e.g., 21.4 ± 0.55 °C). Temperature stability is critical to maintaining the optical alignment.
   2. Install a microscope body that is equipped with an illumination arm for optical fiber connection, a 100X oil-immersion TIRF objective lens, and DIC components.
   3. Place four laser heads (647 nm, 561 nm, 488 nm, and 405 nm, encircled in Figure 1B) and their heat sinks on the optical table, and make sure the emitted laser beams have the same height and are as short as possible to ensure good stability (e.g., 3’’).
      NOTE: If a laser head sits at a shorter height than other lasers, put an aluminum plate with adequate thickness underneath it. Always ensure maximum contact between the heat sinks and the optical table for the best heat dissipation (Figure 1B). The lasers need to be powerful enough for SR imaging. See Table of Materials. It is recommended to have laser clean-up filters in front of diode lasers.
   4. Install a data acquisition card through a peripheral component interconnect (PCI) interface in a workstation and connect lasers with this card. Control the lasers’ ON/OFF behaviors by transistor-transistor logic (TTL) output, and their power adjustment by the analog output of this card (Figure 1C). Install a proper microscopy imaging software (either commercial or homebuilt) to control the data acquisition card, as well as the microscope body.
   5. Mount the mirrors and dichroic beam splitters (590, 525, and 470 long pass filters) to their respective mounts. Use very stable mirror mounts for the mirrors. Use circular splitters with retaining rings to avoid any bending of the dichroic beam splitters (Figure 1D).
   6. Place the mirrors and dichroic beam splitters on the optical table to combine the laser beams (Figure 1B). To achieve the most stable alignment, make the whole arrangement as compact as possible, and use 1’’-thickness optical posts. Arrange the lasers so that the shorter wavelength lasers are closer to the optical fiber coupling (Figure 1B) since short-wavelength lights dissipate more in the air.
   7. Combine laser beams into a single-mode optical fiber. To do so, build a fiber coupler in a cage system through the following steps:
      1. Mount a fiber adapter plate in a z-axis translation mount (Figure 1E, leftmost panel).
      2. Mount an achromatic doublet lens (focal length = 7.5 mm) in a cage plate (Figure 1E, second panel from the left).
      3. Connect the two parts above by extension rods to form a cage. Mount the cage on the optical table with mounting brackets on the 1’’-thick optical posts (Figure 1E, middle panel).
      4. Align the 647-nm laser first with a single-mode optical fiber (FC/APC end to the coupler).
         NOTE: A rough alignment using a multi-mode optical fiber before using the single-mode optical fiber may help the alignment process. Adjust the angles of the mirrors and dichroic beam splitters (Figure 1D, by adjustment knobs), as well as the distance between the achromatic doublet lens and the fiber adapter plate (Figure 1E, by adjusting z-axis translation mount) to gain maximum laser power output through the fiber.
      5. Once the alignment of the first laser is done, temporarily install a pair of irises and align the rest of the lasers one by one (Figure 1F). Check the alignment efficiency with a power meter.
      6. Leave one iris in the front of the adapter plate to reduce the reflections of the lasers.
         NOTE: Strong back reflections can reduce the lifetime of laser sources. Optionally, an optical isolator can be installed in front of each laser head to remove the reflections completely.
   8. Connect the other end of the optical fiber to the illumination arm of the microscope (Figure 1H).
   9. Design and install the “magnification lens (mag lens)” through the following steps:
      NOTE: The lasers can be used for epi-fluorescence imaging of the sample, but the narrow beam size of each laser limits the illuminated area of the sample to an area several times smaller than the actual size of the corresponding camera sensor, especially for the newer cameras (with 18.8 mm in diagonal length compared to the conventional 11.6-mm length). Thus, it is desirable to expand the beam to achieve a larger and flatter illumination of the sample.
      1. Design the mag lens, which can fit into the illumination arm (Figure 1H).
         NOTE: The design of the mag lens depends on where it will be installed. Figure 1H shows an example of the installation in the illumination arm, but it can be installed in any spot after the laser beams are collimated (see Discussion). Design it with a computer-aided design and drafting software.
      2. Place two achromatic lenses, one concave (focal length = f₁), and one convex (focal length = f₂) in the home-made mag lens holder (Figure 2A and 2C), with the distance equal to the sum of their focal lengths, f₁ + f₂ = (-25) + 50 = 25 mm (Figure 2B).
         NOTE: With this choice of focal lengths, the mag lens expands the beam by f₂/|f₁| = 2 folds. The mag lens provides versatility. It can be removed to resume regular illumination without expanding the beams (Figure 2D) or inserted in the reverse direction to focus the laser beam to achieve a stronger excitation intensity.

2. Emission path

NOTE: The emission path is composed of a removable cylindrical lens, a barrier filter wheel, an emission splitter, and an EMCCD camera (Figure 1G). To attain the best point spread function (PSF) of single molecules, the DIC prism is put away from the objective lens.

1. Custom-design the cylindrical lens (3-D lens) cassette, which can fit into the manual DIC analyzer insert slot in the microscope body (Figure 3C).
   NOTE: This design does not compromise the DIC analyzer since an analyzer block can be inserted in the filter turret.
2. Place the 3-D lens of 10 m of focal length in the cassette and insert it into the emission beam path to create the astigmatism effect necessary for extracting the z coordinate of every single molecule¹⁴.
   NOTE: Optionally, the 3-D lens cassette can be placed in or out of the emission path (Figure 3C).
3. Install a multi-band dichroic beam splitter in the filter turret inside of the microscope body.
4. Install emission filters.
   NOTE: The emission filters are chosen depending on the preferred fluorophores. Depending on the imaging module, emission filters placed in different locations are used as described below:
   1. For sequential multi-color epi-fluorescence imaging or SR imaging, use emission filters placed in the barrier filter wheel connected next to the microscope body to minimize the vibration in the microscope body during channel switch (Figure 1G).
   2. For simultaneous multi-color detection (e.g., smFRET experiments), place another filter set in an emission splitter (check step 4 for details).
      NOTE: Usually, a commercial emission splitter has two switchable modes (i.e., “engaged” or “bypass” modes). To separate emission lights chromatically for simultaneous multi-color imaging (“engaged” mode), a filter cube holding two dichroic beam splitters and three emission filters is used (“triple cube,” Figure 4C and 4D). An empty slot in the barrier filter wheel is used in combination with the triple cube. On the other hand, for sequential multi-color imaging, the triple cube is replaced by a cube that just has a mirror inside (“bypass cube,” Figure 4A and 4D).
5. Install the EMCCD camera as the last part of the emission path. Utilize the USB-PCI connection to achieve a fast frame rate.
   NOTE: An EMCCD camera is recommended for the most sensitive single-molecule detection, but an advanced sCMOS camera can be an alternative.

3. Diffraction-limited Imaging with Epi-excitation

1. Adjust the excitation lasers’ incidental angle to epi-mode in the illumination arm.
2. Disengage the 3-D lens if engaged (Figure 3C, right panel).
3. Insert the bypass cube in the emission splitter (Figure 4A and 4E, bottom panel).
4. (Optional) Insert the mag lens for a broadened illumination area (Figure 2D, left panel).
   NOTE: With the use of a mag lens and a 100X oil-immersion objective lens, about 91 x 91 µm² can be illuminated evenly, eliminating the need to use a white light source and multiple filter cubes.
5. Use a microscopy imaging software to take multi-channel, and/or Z-stack, and/or time-lapse images.
   NOTE: There are multiple programs available for microscopy imaging, not only from microscope manufacturers but also from third-party companies or open source developers.

4. Multi-channel Single-molecule Imaging Including smFRET

NOTE: Move to an “empty” position in the barrier filter wheel, so that all the emission with any wavelength can reach to the second set of filters/dichroic beam splitters in the emission splitter.

1. To set up multi-color single-molecule detection of surface-immobilized molecules¹⁵ using TIRF excitation, including smFRET measurement, adjust the excitation lasers’ incidental angle to the TIRF angle. Disengage the mag lens and the 3-D lens.
2. Engage the three-channel mode in the emission splitter (Figure 1G) through the following steps:
   1. Replace the bypass cube with a “calibration cube” that allows light to go through all channels (Figure 4B and 4E).
   2. Turn on the camera under DIC (i.e., no emission filter in the barrier filter wheel) and adjust the aperture of the emission splitter until three fully separated channels appear on the screen.
      NOTE: Conduct this step with the room light lit, to visualize all the channels.
   3. Turn the vertical/horizontal adjustment control knobs on the emission splitter and roughly align the three channels (Figure 4E and 4F).
   4. Turn off the camera and replace the calibration cube with a triple cube (Figure 4C and 4E).
   5. Place a sample with 100-nm multichannel beads on top of the 100X objective lens and focus on the sample.
   6. Turn on the camera and the 488-nm laser, zoom in on one of the bright beads, and finely align the three channels by turning the adjustment control knobs again (Figure 4E and 4G).
      NOTE: 100-nm multichannel beads emit different wavelengths of light upon 488-nm excitation, enabling the three-channel alignment.
3. Turn on the camera and lasers, focus, and find a good position with a reasonable spot density. Adjust the laser power and exposure time to achieve acceptable signal-to-noise and photobleaching levels. Use the microscopy imaging software to take time-lapse images.

5. SR Imaging

NOTE: This is single-molecule detection-based SR microscopy.

1. To set up SR imaging, insert the 3-D lens and remove the mag lens. Set the exposure time of the camera in the appropriate laser channels (e.g., 5 - 60 ms). Determine and manually set the optimal excitation lasers’ incidental angle to be the TIRF angle.
2. Place the sample in SR imaging buffer¹⁶. Allow the buffer to equilibrate for at least 10 min before imaging.
   NOTE: SR imaging buffer expires after roughly 1 h, so make new SR imaging buffer accordingly.
3. Take a DIC image before SR imaging. In order to find the proper objective height for SR, which optimizes the astigmatism effect, use DIC imaging to find the middle plane of the cells. Identify the plane by the height at which the cells transition from “light” to “dark” images and appear to become transparent (Figure 5A, 5B, and 5C). Once the desired focal plane is determined, engage the z-drift correction system (Figure 1A).
4. Conduct SR imaging. Change the 405-nm laser power to maintain a reasonable density of ‘blinking-on’ spots.
   NOTE: While it is possible to change the 405-nm laser manually, it is more convenient to run a programmed data acquisition code to maintain the density of “blinking-on” spots. Here is an example of how it is conducted automatically. The source code is available upon request (Figure 6).
   1. Start the imaging acquisition with 0 W/cm² violet laser power.
   2. Count the number of blinking-on spots in a certain period.
   3. Modulate the violet laser power so that the number of blinking-on spots is kept above a user-defined “counting threshold” in the field of view. Increase the violet laser power when the number of blinking-on spots drops below the counting threshold.
   4. Terminate the acquisition when the number of blinking-on spots drops below the counting threshold using the maximum violet laser power.
      NOTE: The maximum can be set differently depending on the sample brightness, but no higher than 130 W/cm². Depending on the actual goal of the SR imaging, this automatic acquisition code can be manually terminated at any desired point.
5. Check the blinking behavior and PSFs of the spots soon after beginning the acquisition.
   NOTE: If the blinking behavior is not ideal, change the excitation lasers’ incidental angle or replace the imaging buffer. Expect a sampling of “vertical”, “horizontal”, and “diamond” shapes of PSF, representing fluorophores from below, above, and within the focal plane, respectively (Figure 5D). If most spots show either vertical or horizontal PSFs, then the focal plane is off from the center of the cells, so terminate the experiment and adjust the focal plane again. A presence of air bubble in the immersion oil or other local factors can affect the PSFs’ quality, so it may be necessary to replace the oil or change to a different imaging area of the sample.
6. For the data analysis, use either open source (in NIH ImageJ plugins) or commercially available codes to detect centroids of each spot in each imaging frame and extract z-values of each spot from x- and y-widths¹⁴.
   NOTE: In this report, a source code originally developed in one of the earliest single-molecule detection-based SR⁸ was modified for 3-D detection¹⁶ and was used.
7. In the case of two-color imaging, image the fluorophore with the longer excitation wavelength, followed by the one with the shorter excitation wavelength. Run the automated acquisition code similarly to the one described in step 5.4, but with a different imaging laser.
   NOTE: Chromatic aberration should be corrected between images with different fluorophores (e.g., red dye and yellow-green dye). Here are the steps.
   1. Immobilize multiple 100-nm multichannel beads on the glass coverslip, avoiding forming clusters.
   2. Take images of them in different excitation channels.
   3. Extract their (X, Y, Z) coordinates by software (step 5.6).
   4. Plot ΔX_i = X_1i - X_2i and ΔY_i = Y_1i - Y_2i (i is for different beads, and 1 and 2 are different color channels) separately and fit them with proper functions. Save the functions.
      NOTE: Linear functions are sufficient in most cases. Once these functions are determined, this measurement does not have to be repeated each time of imaging.
   5. In the actual two-color SR imaging of a sample of interest, apply the functions to correct (X, Y) chromatic aberration. For z-directional chromatic aberration, conduct it by obtaining ΔZ = Z₁ - Z₂ for multichannel beads or known reference multichannel samples seeded together with the sample of interest.
      NOTE: Unlike (X, Y) chromatic aberration, z-directional chromatic aberration is not well-reproducible in each experiment, mainly due to incomplete z-directional focus maintenance upon channel switching. Thus, it is recommended to conduct the correction each time. ΔZ = Z₁ - Z₂ is mostly independent of (X, Y), so just a few beads or reference samples would be sufficient per each sample area of interest. Plot the constructed final two-color SR images in a 3-D visualization software and check ΔZ manually.

Wyniki

This microscope allows flexible and reproducible switching between different imaging methods. Here we show sample images collected with each imaging module.

Figure 5D demonstrates the PSF of the blinking-on molecule during the SR acquisition. Thousands of such images are reconstructed to generate the final SR image (Figure 5E). Figure 5E sh...

Dyskusje

This hybrid microscope system eliminates the need to purchase multiple microscopes. The total cost for all parts, including the optical table, table installation labor, software, and workstation, is about $230,000. Custom-machined parts, including the mag lens and 3-D lens, cost around $700 (the cost depends on the actual charges at different institutes). Typical commercially available integrated systems for single-molecule detection-based SR microscopy cost more than $300,000 ~ 400,000 and are not readily available for ...

Materiały

Name	Company	Catalog Number	Comments
Nikon Ti-E microscope stand	Nikon	Ti-E
Objective lens	Nikon	100X NA 1.49 CFI HP TIRF
Microscopy imaging software	Nikon	NIS-Elements Advanced Research/HC	HC includes "JOBS" module, the programmed acquisition module being used for SR imaging.
The illumination arm	Nikon	Ti-TIRF-EM Motorized Illuminator Unit M	This arm has a slot for a magnification lens
Analyze block	Nikon	Ti-A	This is installed in the filter turret.
Z-drift correction system	Nikon	PFS	This system is composed by the stepmotor on the objective nosepiece, IR LED, and a detector.
Optical table top	TMC	783-655-02R
Optical table bases	TMC	14-426-35
647 nm laser	Cobolt	90346 (0647-06-01-0120-100)	Modulated Laser Diode 647nm 120mW incl. laser head, CDRH control box, USB cable and PSU (Power Supply Unit)
561 nm laser	Coherent	1280721	OBIS 561nm LS 150mW Laser System
488 nm laser	Cobolt	90308 (0488-06-01-0060-100)	Modulated Laser Diode 488nm 60mW incl. laser head, CDRH control box, USB cable and PSU (Power Supply Unit)
405 nm laser	Crystalaser	DL405-025-O	405 (+/-5)nm, 25mW, Circular , M2 <1.3, Low Noise, CW, TTL up to 20MHz. 2 BNC connectors for TTL & Analog adjust
Heat sink	Cobolt	11658 (HS-03)	Two units, Heat sink without fan HS-03, Heat sink for 647 nm and 488 nm lasers
Heat sink	Coherent	1193289	Obis heat sink with fan, 165 x 50 x 50 mm for the 561 nm laser
CAB-USB-miniUSB	Cobolt	10908	Two units, communication cable for 647 nm and 488 nm lasers
aluminum for height adjustment	McMaster-Carr	9146T35	Multipurpose 6061 Aluminum, Rectangular Bar, 4MM X 40MM, 1' Long for raising 561 nm laser
aluminum for height adjustment	McMaster-Carr	8975K248	Multipurpose 6061 Aluminum, 1-1/4" Thick X 3" Width X 1' Length for raising 405 nm laser
BNC cable	L-com	CC58C-6	RG58C Coaxial Cable, BNC Male / Male, 6.0 ft
BNC adapter	L-com	BA1087	Coaxial Adapter, BNC Bulkhead, Grounded
SMA to BNC Adapter	HOD	SMA-870	Cobolt MLD lasers have SMA interface, so this adapter is used for BNC connection.
SMB to BNC Adapter	Fairview Microwave	FMC1638316-12	SMB Plug to BNC Female Bulkhead Cable RG316 Coax in 12 Inch for Coherent Obis lasers
Data Acquisition Card	National Instruments	PCI-6723	13-Bit, 32 Channels, 800 kS/s Analog Output Device for controlling lasers, DIC LED, and etc
Barrier Filter Wheel controller	Sutter Instrument	Lambda 10-B	Optical Filter Changer
Emission Splitter	Cairn	OptoSplit III
Dichroic beamsplitter	Chroma	T640LPXR-UF2	Dichroic beamsplitter separating red emission from green emission in OptoSplit III
Dichroic beamsplitter	Chroma	T565LPXR-UF2	Dichroic beamsplitter separating green & red emission from blue emission in OptoSplit III
Emission filter	Chroma	ET700/75M	Two units, Emission filter for red emission (like Alexa Fluor 647) in OptoSplit III as well as in the Barrier filter wheel
Emission filter	Chroma	ET595/50M	Two units, Emission filter for yellow/green emission (like Cy3B) in OptoSplit III as well as in the Barrier filter wheel
Emission filter	Chroma	ET525/50M	Two units, Emission filter for blue emission(like Alexa Fluor 488/GFP) in OptoSplit III as well as in the Barrier filter wheel
Emission filter	Semrock	FF02-447/60-25	Emission filter for violet emission (like DAPI/Alexa Fluor 405), installed in the Barrier filter wheel
Dichroic beamsplitter	Chroma	zt405/488/561/647/752rpc-UF3	Multiband dichroic beam splitter for 647, 561, 488, and 405 nm laser excitations inside of the microscope body
DAPI Filter set	Chroma	49000	installed in the microscope body
Nikon laser/TIRF filtercube	Chroma	91032
590 long pass filter	Chroma	T590LPXR-UF1	for combining 647 nm laser and 561 nm laser
525 long pass filter	Chroma	T525LPXR-UF1	for combining already combined 647 nm and 561nm lasers with 488 nm laser
470 long pass filter	Chroma	T470LPXR-UF1	for combining already combined 647 nm, 561 nm and 488 nm lasers with 405 nm laser
Laser clean-up filter (647)	Chroma	zet640/20x	for cleaning up other wavelengths from the 647 nm laser
Laser clean up filter (488)	Semrock	LL01-488-25	for cleaning up other wavelengths from the 488 nm laser
LED light source	Excelitas	X-Cite120LED	used only for DAPI imaging
Mirror mount	Newport	SU100-F3K
Optical posts	Newport	PS-2
Clamping fork	Newport	PS-F
Power Meter	Newport	PMKIT	For measuring laser power
Dichroic beamcombiner mount	Edmund Optics	58-872	C-Mount Kinematic Mount, for holding dichroic beamcombiners in the laser excitation assembly
Retaining ring	Thorlabs	CMRR	used for dichroic beamcombiner mounts
Fiber Adapter Plate	Thorlabs	SM1FC	FC/PC Fiber Adapter Plate with External SM1 (1.035"-40) Thread
Z-axis translational mount	Thorlabs	SM1Z	Z-Axis Translation Mount, 30 mm Cage Compatible
Achromatic Doublet lens	Thorlabs	AC050-008-A-ML	Ø5 mm, Mounted Achromatic Doublets, AR Coated: 400 - 700 nm
Cage Plate	Thorlabs	CP1TM09	30 mm Cage Plate with M9 x 0.5 Internal Threads, 8-32 Tap
Cage Assembly Rod	Thorlabs	ER4	Cage Assembly Rod, 4" Long, Ø6 mm
Cage Mounting Bracket	Thorlabs	CP02B	30 mm Cage Mounting Bracket
Single mode optical fiber	Thorlabs	P5-405BPM-FC-2	Patch Cable, PM, FC/PC to FC/APC, 405 nm, Panda, 2 m
Multi mode optical fiber	Thorlabs	M42L01	Ø50 µm, 0.22 NA, FC/PC-FC/PC Fiber Patch Cable, 1 m
Achromatic Doublet lens (mag lens)	Thorlabs	ACN127-025-A	ACN127-025-A - f=-25.0 mm, Ø1/2" Achromatic Doublet, ARC: 400-700 nm , a concave lens in the "mag lens"
Achromatic Doublet lens (mag lens)	Thorlabs	AC127-050-A	f=50.0 mm, Ø1/2" Achromatic Doublet, ARC: 400-700 nm, a convex lens in the "mag lens"
Retaining ring	Thorlabs	SM05PRR	SM05 Plastic Retaining Ring for Ø1/2" Lens Tubes and Mounts, for "mag lens"
Nylon-tipped screw	Thorlabs	SS3MN6	M3 x 0.5 Nylon-Tipped Setscrew, 6 mm Long, for holding "3D lens"
3D lens	CVI Laser Optics	RCX-25.4-50.8-5000.0-C-415-700	f=10 m, rectangular cylindrical lens
EMCCD camera	Andor	iXon Ultra 888
100 nm multichannel beads	Thermo	T7279, TetraSpeck microspheres
red dye	Thermo	Alexa Fluor 647
yellow-green dye	GE Healthcare	Cy3
green dye	GE Healthcare	Cy3B
blue dye	Thermo	Alexa Fluor 488