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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

This protocol details the construction and operation of a real-time 3D single particle tracking microscope capable of tracking nanoscale fluorescent probes at high diffusive speeds and low photon count rates.

Abstract

Real-time three-dimensional single particle tracking (RT-3D-SPT) has the potential to shed light on fast, 3D processes in cellular systems. Although various RT-3D-SPT methods have been put forward in recent years, tracking high speed 3D diffusing particles at low photon count rates remains a challenge. Moreover, RT-3D-SPT setups are generally complex and difficult to implement, limiting their widespread application to biological problems. This protocol presents a RT-3D-SPT system named 3D Dynamic Photon Localization Tracking (3D-DyPLoT), which can track particles with high diffusive speed (up to 20 µm2/s) at low photon count rates (down to 10 kHz). 3D-DyPLoT employs a 2D electro-optic deflector (2D-EOD) and a tunable acoustic gradient (TAG) lens to drive a single focused laser spot dynamically in 3D. Combined with an optimized position estimation algorithm, 3D-DyPLoT can lock onto single particles with high tracking speed and high localization precision. Owing to the single excitation and single detection path layout, 3D-DyPLoT is robust and easy to set up. This protocol discusses how to build 3D-DyPLoT step by step. First, the optical layout is described. Next, the system is calibrated and optimized by raster scanning a 190 nm fluorescent bead with the piezoelectric nanopositioner. Finally, to demonstrate real-time 3D tracking ability, 110 nm fluorescent beads are tracked in water.

Introduction

The emergence of advanced imaging techniques has opened a window to see ever more detailed structure of cellular phenomena, all the way down to the molecular level. Methods such as stochastic optical reconstruction microscopy (STORM)1,2,3, photo-activated localization microscopy (PALM)4,5,6,7, structured illumination microscopy (SIM)8,9,10,11, and stimulated emission depletion microscopy (STED)12,13,14 have gone far beyond the diffraction limit to deliver unprecedented detail into the structure and function of live cells. However, full understanding of how these systems behave requires dynamic information as well as structural information. The super-resolution methods listed above involve a trade-off between spatial resolution and temporal resolution, limiting the temporal precision with which dynamic processes can be probed. A method which provides both high spatial precision and temporal resolution is RT-3D-SPT15,16,17,18,19,20,21,22,23,24,25,26,27,28,29. Here, we draw a distinction between traditional 3D-SPT30 and RT-3D-SPT. Traditional 3D-SPT simply requires a time series of three-dimensional image data (which can be acquired either using a confocal microscope or an epifluorescence microscope given the right configuration). In traditional 3D-SPT, the coordinates of the particle are determined after data collection by locating the particle in each image stack and concatenating the locations in successive volumes to create a trajectory. For these methods, the ultimate temporal resolution is determined by the volumetric imaging rate. For confocal microscopes, this is easily on the scale of seconds to tens of seconds. For epifluorescence methods, wherein the optical path is manipulated so that the axial location information can be extracted, the temporal resolution is limited by the camera exposure or readout time. These epifluorescent methods are limited in the range over which axial information can be collected, though recent progress in Fourier plane phase masks design and adaptive optics is extending these ranges to 10 µm or more31,32,33,34.

In contrast, RT-3D-SPT does not rely on acquiring a 3D image stack and finding particles after the fact. Instead, real-time location information is extracted via single point detectors and feedback is applied to effectively "lock" the particle in the focal volume of the objective lens through the use of a highspeed piezoelectric stage. This allows continuous measurement of the particle's position limited only by how many photons can be collected. Moreover, this method enables spectral interrogation of the particle as it moves over long ranges. RT-3D-SPT in effect works akin to a force-free optical trap for nanoscale objects, wherein the particle is continuously probed and measured in real-time without the need for large laser powers or optical forces. Given that RT-3D-SPT provides a means for continuous interrogation of fast diffusive objects (up to 20 µm2/s)25,29 in three dimensions at low photon count rates20,29,35, it should provide a window into fast or transient biological processes such as intracellular cargo transport, ligand-receptor binding, and the extracellular dynamics of single virions. However, to this point, the application of RT-3D-SPT has been limited to the handful of groups working to advance this technology.

One barrier is the complexity of the optical layout required by RT-3D-SPT methods, which are varied. For most methods, the optical feedback is provided by a piezoelectric stage. As the particle makes small movements in X, Y, or Z, readouts from single point detectors are converted to error functions and fed at high-speed to a piezoelectric nanopositioner, which in turn moves the sample to counteract the particle's motion, effectively locking it in place relative to the objective lens. To measure small positional movements in X, Y, and Z, either multiple detectors (4 or 5 depending on the implementation)15,18,21 or multiple excitation spots (2 - 4, the lower of which can be applied if a lock-in amplifier is used to extract X and Y position using a rotating laser spot)25,28 are applied. The overlap of these multiple detection and emission spots make the systems difficult to align and maintain.

Herein, we present a high-speed target-locked 3D-SPT method with a simplified optical design called 3D-DyPLoT29. 3D-DyPLoT uses a 2D-EOD and a TAG lens36,37,38 to dynamically move a focused laser spot through the objective focal volume at a high rate (50 kHz XY, 70 kHz Z). Combining the laser focus position and the photon arrival time enables the particle's 3D position to be rapidly obtained even at low photon count rates. The 2D-EOD drives the laser focus in a knight's tour pattern39 with a square size of 1 x 1 µm in the X-Y plane and the TAG lens moves the laser focus in axial direction with a range of 2 - 4 µm. The 3D particle position is obtained with an optimized position estimation algorithm29,40 in 3D. The control of the 3D dynamically moving laser spot, photon counting from the avalanche photodiode (APD), real-time particle position calculation, piezoelectric stage feedback, and data recording are performed on a field programmable gate array (FPGA). In this protocol, we describe how to build a 3D-DyPLoT microscope step-by-step, including optical alignment, calibration with fixed particles, and finally free particle tracking. As a demonstration, 110 nm fluorescent beads were tracked continuously in water for minutes at a time.

The method described herein is an ideal choice for any application where it is desired to continuously monitor a fast-moving fluorescent probe at low light levels, including viruses, nanoparticles, and vesicles such as endosomes. In contrast to previous methods, there is only a single excitation and single detection pathway, making alignment and maintenance straightforward. Furthermore, the large detection area enables this microscope to easily pick up quickly diffusing particles, while the ability to track at low signal levels (down to 10 kHz) makes this method ideal for low-light applications29.

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Protocol

1. Setup Layout and Alignment

  1. Installation and collimation of the tracking excitation laser
    1. Affix the laser to the optical table using a home-built mount. The mount is a simple aluminum plate with mounting holes for the laser head and the optical table. The laser should be firmly attached to a metal mount for stability and heat dissipation. For this work, use a 488 nm solid state laser for illumination (Figure 1), though the wavelength can be selected to suit a particular fluorophore or experiment. A critical factor is that the laser wavelength fit the 2D-EOD working range, which is determined primarily by the wave plate between the two deflectors (Figure 1: W2). The 488 nm laser can effectively excite the green or yellow fluorescent protein (GFP/YFP), which are common fluorescent tags in live cell experiments.
    2. Adjust the laser beam height and direction using a pair of dielectric mirrors (Figure 1: M). Make sure the laser beam is parallel to the optical table and adjust it to a suitable height.
      NOTE: The height should be chosen based on the microscope platform. This height should be maintained for all subsequent mirrors used in this protocol, though it will not be explicitly stated.
    3. Collimate the beam with a pair of lenses (Figure 1: L1 and L2). The focal lengths of lenses should be carefully chosen to avoid the laser spot being clipped by the aperture of the 2D-EOD. Clipping is evident by a change in the laser beam shape after passing through the 2D-EOD. The quality of collimation here is very important because aberrations will be amplified by subsequent lenses and any divergence will deteriorate the performance of the 2D-EOD deflection. Ideally, collimate the beam by focusing the beam to a point at least 20 m away.
  2. Place a pinhole (Figure 1: PH) of appropriate size at the focus of the first collimation lens (Figure 1: L1).
    1. Choose an appropriately sized pinhole. The pinhole size can be chosen based on the following calculation:
      figure-protocol-2351
      Where λ is the wavelength of laser, f is the focal length of the first lens, and D is the input beam diameter.
    2. Mount the pinhole on a 3D translation stage so that it can be placed precisely at the focus of the first collimation lens.
    3. Adjust the position of the pinhole. Place a laser power meter after the pinhole and adjust the pinhole position in XYZ to maximize the power meter readout. If a diffraction ring is observed, block it with an iris placed before the 2D-EOD.
  3. Installation of Glan-Thompson polarizer
    1. Put the Glan-Thompson polarizer (Figure 1: GP) after the second collimation lens (Figure 1: L2) to clean the polarization of the laser beam. Rotate the polarizer to find maximum transmission with the power meter.
  4. Installation of EODs
    1. Use 2 EODs (Figure 1: EOD1 & EOD2) to deflect the laser in the X and Y directions. Maximize the laser transmission by adjusting the yaw, pitch, and height of each EOD.
    2. Align the two EODs with respect to each other using the alignment marker provided by the manufacturer on the side of each deflector.
    3. Apply a test pattern to the controller of the 2D-EOD to inspect the output of the EOD. This can either be the knight's tour (Figure 2) via the FPGA described below or a simple sine wave provided by a function generator. For the best performance, the deflection by each deflector should be parallel to either the X or Y direction of the piezo nanopositioner. This direction is dictated by the angular direction of the deflectors about the transmission axis. To rotate the deflection direction without moving the 2D-EOD, place a dove prism after the 2D-EOD to align the deflection axes to the piezo nanopositioner axes.
  5. Installation of half-wave plate
    1. Place a half-wave plate (Figure 1: W1) between the Glan-Thompson polarizer (Figure 1: GP) and the 2D-EOD to align the incoming laser beam polarization with the deflection axes of the 2D-EOD. Place a long focal lens (300 mm) after the 2D-EOD and place a sCMOS camera at the laser focus. Next, turn on the 2D-EOD to generate the knight's tour described below (Figure 2). Rotate the half-wave plate until an evenly square laser distribution is observed on the camera.
  6. Installation of a beam expending lens pair and relay lens pair
    1. Place a pair of lenses (Figure 1: L3 and L4) after the 2D-EOD to expand the beam to fill the back aperture of the objective lens (Figure 1: OL) and another pair of lenses (Figure 1: L5 and L6) to relay the focal plane to the microscope objective. Be sure to leave enough room between these two pairs of lenses as the TAG lens will be installed between them in a later step.
  7. Install the dichroic filter, 10/90 beam splitter, focusing lens, APD, objective lens, and fluorescence emission filter to complete the detection pathway.
    1. Install the dichroic filter (Figure 1: DC) to reflect the laser towards the objective lens. Align the laser to go straight through the center of the objective lens using two irises on the top and bottom of a long lens tube.
    2. Install a 10/90 beam splitter (Figure 1: BS) by which 10% of light is reflected for imaging by an sCMOS camera (described below) and 90% of passes through to the APD for tracking.
    3. Align the APD. Focus the laser onto a coverslip by the objective lens. Use the reflection from the coverslip to align all downstream elements by putting a coverslip at the objective focus and checking the intensity of the APD readout. After the beam splitter, install the APD focusing lens (Figure 1: L7) followed by the APD on a 3D translation stage. Position the lens such that the laser reflection from the objective goes through the center of the lens. The APD position can be optimized by adjusting the 3D position to maximize the intensity from the laser reflection on the coverslip. The APD position is at the optimal position when a move of the detector position in any direction decreases the intensity.
    4. Install a longpass fluorescence emission filter (Figure 1: F) before the beam splitter to remove the reflected and scattered light.
  8. Installation of the TAG lens
    1. Place the TAG lens between the two pairs of lenses (between Figure 1: L4 and L5) as mentioned before and shown in Figure 1. Adjust the yaw, pitch, height, and lateral position such that the beam passes perpendicularly through the center of TAG lens.

2. Sample Preparation.

  1. Fixed particle preparation
    1. Dilute 190 nm fluorescent beads to ~ 5 × 108 beads/mL in PBS. Add 400 µL beads solution onto a coverslip and mount on the sample holder of the piezoelectric stage (the volume of beads will depend on the sample holder; the diameter of the sample holder chamber used here is 18 mm). For the beads used here, PBS causes the particles to deposit on the coverslip.
  2. Free moving particle preparation
    1. Dilute 110 nm fluorescent beads to ~ 5 × 108 beads/mL with DI water. Add 400 µL beads solution onto a coverslip and mount onto the sample holder of the piezoelectric stage.

3. Optimize Tracking Parameters

  1. Raster scan fixed particles
    1. Put a fixed particle sample on the microscope.
    2. Turn on the laser, micropositioner controller, piezo nanopositioner controller, TAG lens controller, and EOD controller. Note that the order is not critical. Run the piezo nanopositioner in closed loop.
    3. Raster scan the sample in XYZ using a custom scanning software program (Figure S3, software available upon request from the authors), which drives the 2D-EOD in a knight's tour (Figure 2), collects counts from the APD, drives the piezo nanopositioner, and performs position calculation (Figure S2). The step size is 40 nm and the scanning range is 2 µm.
      1. To place the coverslip at the focus of the objective, remove the fluorescence emission filter and use the reflection of the laser from the coverslip to maximize the signal intensity as a function of the Z position of the micropositioner. After placing the sample at the focal plane, place back the fluorescence emission filter.
      2. Open the scanning program. Set the scanning range and step size by typing the number in the 'start', 'finish', and 'step'. First set a large scanning range and step size to locate a particle (e.g., 10 x 10 µm range with 200 nm step size). After finding the particle, shrink the scanning range and decrease step size (e.g., 2 x 2 µm range with 100 nm step size). Click the 'scan' button to perform a 3D raster scan to find the fluorescent focus.
  2. TAG lens settings (see also Figure 3)
    1. Click on the TAG lens control software. Click 'connect', 'power on' sequentially.
    2. To change the output phase of the trigger signals, it is necessary to change the output trigger mode. First change the mode from 'RGB' to 'Multiplane' and then change it back. Now the phase can be changed (Figure 3b).
    3. Set the output phase to be 0 °, 90 °, and 270 °. While any three phases which cover the phase space can be used, these three are found to work well empirically (Figure 3c).
    4. Select the 68,500 Hz frequency setting.
    5. To find the optimal frequency it is often necessary to change the frequency search range. Click 'advanced', 'setting'. Change the 'Max. Freq(Hz)' of column 0 from 70,000 Hz to 71,500 Hz. This can be adjusted to whatever frequency range is appropriate. Click 'Save Calibration', 'Exit Calibration' (Figure 3d).
    6. To make the new calibration effective, switch the resonance to another frequency (for example, 189,150 Hz) and then switched back to the 68,500 Hz frequency setting (Figure 3e).
    7. Change the amplitude gradually to 35%. Click 'Lock Resonance'. After the frequency is locked, click 'Unlock Resonance' (Figure 3e). The TAG lens is ready to use now.
    8. To calibrate, change the parameters that are used in the estimation to make the estimated position equal to the real position, as shown in Figure 4e, f. Input the scanning range of x, y, and z and then click the 'scan' button to scan the sample in XYZ to move the particle through the moving laser spot. The position of the particle as it is scanned through the laser focus should agree with the estimated particle position determined by the position estimation loop (Figure S1). The relationship between the estimated position and real position (Figure 4d) can be extracted from estimated position images (Figure 4f). If the positions do not agree, adjust the values of laser position (ck) used in the position estimation loop (Figure S1).
    9. Align TAG lens
      1. When the TAG lens controller is off, the raster scanned particle images (described below) taken before installing TAG lens and after installing TAG lens should look identical. Next, perform fluorescent particle Z stack scanning by moving the objective with the z nanopositioner to tune the position and angle of TAG lens (Figure 4). Tune the position of the TAG lens until there is no drift in the particle images in XY plane along the Z direction, which is achieved when there is no change in the XY position of the particle as a function of the Z position (Figure 4d). The tracking system is ready to use after alignment of the TAG lens.
  3. Installation of sCMOS camera for particle monitoring
    1. Install a sCMOS camera to visualize particles while they are being tracked. Install the sCMOS only after the all the components of the tracking system have been installed and optimized. Load a fixed fluorescent particle sample onto the microscope and run the tracking program to lock one particle in the objective focal volume. Then, install the 100 mm focal length lens (Figure 1: L8) and sCMOS. Adjust the sCMOS position so that the image of the particle is focused to the smallest and brightest spot.

4. Real-time 3D Tracking of Freely Diffusing Nanoparticles

  1. Put the 110 nm free moving particle sample on the microscope.
  2. Turn on laser, micro-positioner controller, piezo nanopositioner controller, TAG lens controller, and EOD controller. Run the piezo nanopositioner in open loop. Set the TAG lens software according to step 3.2.
  3. Open and run the tracking software (available upon request from the authors). Set the position estimation parameters to their optimized values, which were found in Section 3.
  4. To place the coverslip at the focus of the objective, remove the fluorescence emission filter and use the reflection of the laser from the coverslip to maximize the signal intensity as a function of the Z position of the micropositioner. After finding the coverslip, increase the focus position 15 µm to focus the laser away from the coverslip and into the solution. Place back the fluorescence emission filter.
  5. Set the integral control constants. The integral control constants can be set starting at a low value and slowly increasing until oscillations can be seen in the particle's position readouts. Once oscillations are observed, set the integral control constants to 80% of the value causing the oscillations. Typical values will be around 0.012 for the XY 'integral gain' and 0.004 for the Z 'integral gain'.
  6. Set the tracking thresholds in 'Track Threshold' and 'Track Minimum' and start the tracking experiment by clicking 'Search' and 'Auto Track'. The threshold for terminating tracking is set slightly higher than the background level and the threshold for triggering tracking is about two times higher than the background. Here, the threshold for triggering tracking is 3 kHz and the threshold for ending the trajectory is 1.5 kHz.

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Results

Fixed particle scanning (Figure 4) and freely diffusing 110 nm fluorescent particle tracking (Figure 5) were performed following the protocol above. The particle scanning was performed by moving the piezoelectric nanopositioner and bin photons while simultaneously calculating the particle's estimated position at each point in the scan. The scanning image shows a square of even intensity (Figure 4a

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Discussion

Although many varieties of 3D single particle tracking methods have emerged in recent years, robust real-time tracking of high speed 3D diffusion at low photon count rates with a simple setup is still a challenge, which limits its application to important biological problems. The 3D-DyPLoT method described in this protocol addresses these challenges in a several ways. First, the excitation and detection pathways are simplified greatly compared to other implementations making alignment simple and robust. Secondly, the mov...

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Disclosures

The authors declare no competing financial interests.

Acknowledgements

This work was supported by the National Institute of General Medical Sciences of the National Institutes of Health under award number R35GM124868 and by Duke University.

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Materials

NameCompanyCatalog NumberComments
2D Electro-optic Deflector ConOpticsM310A2 required
Power supply for EODConOptics412Converts FPGA ouput to high voltage for EOD
TAG  LensTAG OpticsTAG 2.0Used to deflect laser along axial direction
XY piezoelectric nanopositionerMadCity LabsNano-PDQ275HSUsed for moving the sample to lock the particle in the objective focal volume in 
Z piezoelectric nanoposiitonerMadCity LabsNano-OP65HSUsed to move the objective lens to follow the diffusing particle
MicropositionerMadCity LabsMicroDriveUsed to coarsely position sample and evaluate
Objective LensZeissPlanApoHigh numerical aperture required for best sensitivity. 100X, 1.49 NA, M27, Zeiss
sCMOS cameraPCOpco.edge 4.2Used to monitor the particle's position
APDExcelitasSPCM-ARQH-15Lower dark counts beneficial
Field programmable gate arrayNational InstrumentsNI-7852r
SoftwareNational InstrumentsLabVIEW
Tracking excitation laserJDSUFCD488-30
LensThorLabsAC254-150-A-MLL1
LensThorLabsAC254-200-A-MLL2
PinholeThorLabsP75SPH
Glan-Thompson PolarizerThorLabsGTH5-AGT
Half-wave plateThorLabsWPH05M-488WP
LensThorLabsAC254-75-A-MLL3
LensThorLabsAC254-250-A-MLL4
LensThorLabsAC254-200-A-MLL5
LensThorLabsAC254-200-A-MLL6
Dichroic MirrorChromaZT405/488/561/640rpcDC
Fluorescence Emission FilterChromaD535/40mF
10/90 beamsplitterChroma21012BS
PBSSigmaD8537
190 nm fluorescent nanoparticlesBangs laboratoriesFC02F/9942
110 nm fluorescent nanoparticlesBangs laboratoriesFC02F/10617
CoverslipFisher Scientific12-545A
PowermeterThorlabsPM100D
CMOSThorlabsDCC1545M
IrisThorlabsSM1D12D
MicroscopeMad City LabsRM21

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