Name: Microfluidic Imaging Flow Cytometry by Asymmetric-detection Time-stretch Optical Microscopy (ATOM) (Video) | JoVE Journal
Uploaded: 2017-06-28T15:00:00.000+00:00
Duration: 7 min 19 s
Description: Learn to implement ATOM for label-free high contrast single-cell imaging in ultrafast microfluidic flow. This method enables imaging at 100,000 cells per second with enhanced contrast for unlabeled cells.

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5 minutes

We thank Mr. P. Yeung for preparing the MCF-7 for us. This work was partially supported by grants from the Research Grant Council of the Hong Kong Special Administration Region, China (Project no. 17259316, 17207715, 17207714, 17205215, and HKU 720112E), the Innovation and Technology Support Programme (ITS/090/14), and the University Development Fund of HKU.

1. Sample Preparation
<ol>
	<li>Sample preparation (adherent cells; MCF-7 cells)
	<ol>
		<li>Take out the cell culture dish from the incubator and drain the culture medium.</li>
		<li>Rinse the cells on a dish with 1x phosphate-buffered saline (PBS) to remove excessive culture medium.</li>
		<li>Add 3 mL of a solution of 0.25% trypsin to the culture dish (diameter of 100 mm) and put it in a 37 °C, 5% CO2 incubator for 4 min. 
		NOTE: The trypsin dissolves the adhesive protein of the cells so that they will detach from the culture dish.</li>
		<li>Check to see if all the cells have detached from the culture dish using a light microscope (10X objective). If not, gently shake the cell culture dish.</li>
		<li>Add 4 mL of the standard culture medium (formulated with 89% Dulbecco's Modified Eagle Medium (DMEM), 10% fetal bovine serum (FBS), and 1% penicillin-streptomycin (PS)) to stop the action of the trypsin.</li>
		<li>Transfer the whole mixture (cells in trypsin solution and culture medium) to a centrifuge tube and centrifuge for 5 min at 200 x g.</li>
		<li>Remove all the fluid and re-suspend the cells in 1 mL of 1x PBS (pH value: 7.4, pre-heated to 37 °C).</li>
	</ol>
	</li>
	<li>Sample preparation (cultured micro-algae)
	<ol>
		<li>Transfer the cultured micro-algae and the culture medium (e.g., sea water, agar, or fresh water) to new glass culture tubes (~15 mL) in a volume ratio (micro-algae to culture medium) of 1:5 to establish a new sub-culture medium.</li>
		<li>Place the glass culture tubes in a dust-proof chamber under constant illumination by artificial lighting from fluorescent bulbs according to a light/dark (LD) cycle (usually LD 16:8) for 72 - 120 h before the experiment.</li>
		<li>Transfer the sub-cultured samples to a centrifuge tube and mix well.</li>
	</ol>
	</li>
</ol>
2. ATOM System Setup
<img alt="Pulsed laser spectroscopy diagram showing time-stretch, spectral-encoding, asymmetric detection setup." class="xfigimg" src="/files/ftp_upload/55840/55840fig1.jpg" data-altupdatedat="1788250015000" data-alt="Figure 1"> 
Figure 1: Schematic of an ATOM System. A broadband pulsed laser is employed to deliver ultrafast pulses to (a) a time-stretch module and (b) an in-line optical amplifier module. The time-stretch module generates a train of temporal waveforms, each of which is the replica of the wavelength spectrum of the laser source, (i.e., wavelength-to-time mapping). The amplifier module is used to for pulse-stretching (dispersive) loss compensation. The stretched pulse is then (c) spatially dispersed by a diffraction grating, forming a 1D spectral shower illumination in which individual wavelength components are relayed by a relay lens pair and focused by the objective lens onto different positions on the flowing cell inside the (d) microfluidic chip. This is the process of spectral-encoding. The spectrally encoded light will again pass the cell through another objective lens and a mirror, returning to the diffraction grating and recombining as a spatially undispersed pulsed beam. This image-encoded pulsed beam is then (e) split into two paths, such that both beams are partially blocked with (f) the knife edge, but from opposite directions, before being coupled into the fibers. These two beams represent the two (opposite) encoded phase-gradient contrasts of the final image. For the simultaneous detection of both signal contrasts, one of the signals undergoes (g) a time-delay line, such that the two signals are multiplexed (interleaved) in time. A high-speed photodetector and real-time oscilloscope are used for data acquisition. <a href="http://cloudfront.jove.com/files/ftp_upload/55840/55840fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Time stretch module
	<ol>
		<li>Employ a broadband femtosecond (fs) or picosecond (ps) pulsed laser source with a recommended center wavelength in the near infrared (NIR) range, 800-1,500 nm. 
		NOTE: The typical required pulse width can range from sub-100 fs to a few ps. Detailed requirements of the pulsed laser sources can be seen in References 4 and 5. Important metrics are highlighted in Table 1.
		<ol>
			<li>Ensure that the laser source has a high repetition rate, which should be in the megahertz regime (e.g., tens of MHz) to ensure ultrafast imaging in ATOM. Also, set the peak output laser power well below the damage power threshold of the fiber cavity, approximately 1 kW.</li>
			<li>Ensure good shot-to-shot temporal and spectral stability of the pulsed laser source. 
			NOTE: Typical tolerance in the spectral amplitude fluctuation should be kept within 1.2%19,20,21, as achieved by the fiber mode-locked laser in this setup. 
			NOTE: The optical bandwidth of the laser source is expected to be 10-100 nm, which is essential to guarantee sufficient imaging field-of-view (FOV) in single-cell imaging21.</li>
		</ol>
		</li>
		<li>Through a fiber collimator, couple the laser pulsed beam to a single-mode dispersive optical fiber, in which the pulses are stretched via group velocity dispersion (GVD) (Figure 1a). 
		NOTE: This is the process after which the spectrum of each pulse is mapped onto time as a wavelength-swept waveform (i.e., wavelength-to-time mapping). 
		NOTE: The total required GVD should be sufficient to ensure that the overall ATOM image resolution is not affected by the wavelength-to-time mapping process (See the Discussion). Typically, in the NIR range, the GVD should be well beyond 0.1 ns/nm. For instance, a single-mode fiber employed in this current setup provides a total GVD of 0.38 ns/nm around the wavelength of 1,060 nm (a total fiber length of 10 km).</li>
	</ol>
	</li>
	<li>Spectral-encoding module
	<ol>
		<li>Construct an optical microscope system to perform the spectrally encoded imaging of the cells flowing along the microfluidic channel, as illustrated in Figure 1. 
		NOTE: The key components of this optical microscope include: (1) a diffraction grating, a telescopic relay-lens module (RL1 and RL2 in Figure 1), and two objective lenses (Obj1 and Obj2 in Figure 1).
		<ol>
			<li>First, illuminate the time-stretched and collimated beam onto a diffraction grating (transmission-type grating is employed in this setup) to generate a spectral shower (Figure 1c). 
			NOTE: The power of the diffracted beam (i.e., diffraction efficiency) can be maximized by adjusting the grating orientation closed to the Littrow configuration.</li>
			<li>Configure the two relay lenses (RL1 and RL2) in a 4-f imaging system (i.e., place the diffraction grating on the focal plane of RL1 and separate RL1 and RL2 by a distance equal to the sum of their focal lengths. The spectral shower will then be imaged onto the back focal plane of the objective lens Obj1).</li>
			<li>Carefully align the spectral shower to fill the back aperture of the Obj1, such that the spectral shower can be projected and focused onto the image plane of the microscope. 
			NOTE: Here, the NA of the objective lens (Obj1 in Figure 1) is 0.75.</li>
			<li>Place another objective lens (Obj2 in Figure 1) with a similar NA and a plane mirror at the back aperture of this objective lens (Obj2), such that the spectral shower beam can be aligned to double-pass the image plane and return to the diffraction grating.</li>
			<li>Adjust the pair of objective lenses (Obj1 and Obj2) such that their focal planes overlap with each other. Check to see if the spectral shower double-passes the image plane at the same location and returns to the grating following the same path. If not, perform further alignment and tuning of the optical system. 
			NOTE: The returned light should pass through an additional beam-splitter, such that the light can be transmitted to an asymmetric detection module.</li>
			<li>Adjust the optical amplifier gain at a suitable level, such that the resultant signal can be detected by the photodetector with a good SNR, which is typically at &gt;10 dB.</li>
			<li>Place and adjust the position of the microfluidic chip on the sample platform and make sure that the spectral shower, and thus the imaging area, is placed across the microfluidic channel (Figure 1d). 
			NOTE: The spectral shower is illuminated orthogonally to the flow direction of fluidics inside the microfluidic chip, such that the flowing motion automatically performs two-dimensional (2D) scans.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Asymmetric detection module
	<ol>
		<li>Place an additional beam-splitter to separate the imaged-encoded beam into two (Figure 1e). 
		NOTE: Each beam replica is partially blocked by a knife edge.</li>
		<li>Measure and record the optical power of the beam before the beam block. Then, manually position the knife edges (mounted on a linear translation stage) such that they roughly block half of the beam (by visual inspection). Afterward, use the optical power meter to monitor the beam power while fine-tuning the position of the knife edges by translating them across the beam. 
		NOTE: It is suggested that the optimal position is where the power is decreased by ~50% of the original value (i.e., the unblocked case). This is the condition that provides the best combination of image signal strength and image contrast enhancement.</li>
		<li>Repeat step 2.3.2 for another beam replica. Note that the orientation of the partial beam block for one beam should be opposite with respect to the other beam (Figure 1f).</li>
		<li>Couple the two partially-blocked beams into two separate, single-mode fiber arms through two-fiber collimators. 
		NOTE: One of the arms has an extra length of fiber, serving as a fiber-based delay-line, to introduce a time-delay with respect to the other replica (without the delay-line) (Figure 1g). Both are directed to a single fiber by a fiber coupler before photodetection. The time-delay should be long enough to temporally separate the two replicas and short enough to avoid temporal overlap with the next waveform (i.e., the two replicas are always time-interleaved and time-multiplexed in a single fiber prior to detection (Figure 2a)).</li>
		<li>Capture and digitize the detected optical signals with the real-time oscilloscope. Note that the bandwidth and thus the sampling rate of the oscilloscope should be sufficiently high to ensure that they do not influence the final image resolution (See the Discussion). 
		NOTE: Here, with the GVD of 0.38 ns/nm, a backend acquisition bandwidth and sampling rate of &gt;20 GHz and &gt;40 GSa/s, respectively, are required.</li>
	</ol>
	</li>
</ol>
3. Experimental Procedures
<ol>
	<li>Specimen loading
	<ol>
		<li>Perform cell counting under a conventional phase contrast microscope in a standard hemocytometer.</li>
		<li>Adjust the cell density by diluting with 1x PBS (with spring water for micro-algae) and mix well with a pipette. 
		NOTE: The suggested concentration is from 105 to 106 cells/mL.</li>
		<li>Mount the microfluidic chip onto the sample platform of the optical imaging system. 
		NOTE: The microfluidic chip is primarily made of polydimethylsiloxane (PDMS) and is fabricated using the standard replica molding method. The microfluidic channel is designed with an asymmetric curved channel to generate the inertial flow focusing effect, such that the cells can flow in single file in the imaging section (with a dimension of 80 µm x 80 µm (height x width)) at high speed.</li>
		<li>Transfer the density-adjusted cell solution to a 10 mL syringe.</li>
		<li>Connect the syringe to the inlet of the microfluidic chip and a centrifugal tube to the outlet of the microfluidic chip for disposal collection.</li>
		<li>Mount the syringe onto a syringe pump and set a suitable flow rate to give desirable throughput and to avoid imposing excessive shear force between the cells and the channel. 
		NOTE: The linear speed of the samples should be within 0.5 and 10 m/s. Note that before the actual image recording, it is often necessary to further fine-tune the system, in terms of maximizing the image signal strength and optimizing the image focusing by repeating steps 2.2.1.4-2.2.1.6.</li>
	</ol>
	</li>
</ol>
4. Data Acquisition
<ol>
	<li>Set a suitable number of data points to be saved for each experiment. The number of data points to be saved depends on the size and flow rate of the samples. 
	NOTE: Typically, it is set to 8-16 million sample points under a sampling rate of 80 GSa/s.</li>
	<li>Acquire and save the data in a batch mode using the oscilloscope.</li>
</ol>
5. Backend Processing
<img alt="Absorption contrast and phase-gradient imaging diagram; data analysis of optical microscopy setup." class="xfigimg" src="/files/ftp_upload/55840/55840fig2.jpg" data-altupdatedat="1788250015000" data-alt="Figure 2"> 
Figure 2: Reconstruction of ATOM Images from the Line-scan Time Trace. To reconstruct the ATOM images with the absorption contrast and differential (enhanced) phase-gradient contrast, two sets of time-stretched pulses (see the purple and green pulses) are extracted from (a) the time-multiplexed temporal waveform trace and are then digitally segmented and stacked to form (b) two 2D images showing the two opposite phase-gradient contrasts. Note that a shearing operation is needed to form (c) the distortion-free images by compensating for the sub-index shift due to the round-off error in the laser repetition rate estimation. By subtracting the (d) raw line-scan from the spectral intensity envelope of the laser source, the background of the images can be removed. (g) An absorption contrast image can be obtained by a pixel-by-pixel intensity summation of images (e) and (f), whereas (h) a differential phase-gradient contrast image can be obtained from the subtraction of images (e) and (f). <a href="http://cloudfront.jove.com/files/ftp_upload/55840/55840fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol>
	<li>Transfer the data from the oscilloscope to the computers for offline image reconstruction. 
	NOTE: Here, the data are stored in a portable hard disk and transferred manually between the oscilloscope and the processing computers. The capacity of the hard disk should be more than 500 GB for each experiment.
	<ol>
		<li>Use the key image reconstruction routine to digitally stack the line-scans to form a 2D image (Figure 2). 
		NOTE: The custom program (in MATLAB) then produces two asymmetric detection images by separating the two time-multiplexed data sets (purple and green waveforms in Figure 2).
		<ol>
			<li>Obtain a differential phase-gradient contrast image by subtracting the intensities of the two asymmetric detection images; an absorption contrast image can be obtained by adding the two asymmetric detection images. See selected images of MCF-7 and micro-algae in Figure 3. Note that the background profiles of individual images should first be eliminated, and their intensities should be normalized, prior to the image summation and subtraction operations.</li>
		</ol>
		</li>
		<li>Generate a library of parameters derived from the images, such as cell volume, circularity, and absorption density, etc. for further analysis.</li>
	</ol>
	</li>
	<li>Input the library of data into the data visualization platform (Figure 4).
	<ol>
		<li>Load the library of parameters and the corresponding reconstructed images to the visualization interface platform.</li>
		<li>Set the axes to be the parameters of interest. 
		NOTE: The parameters are problem-specific and are defined by the user. They are derived from the ATOM images by the MATLAB program. Here, the optical absorption density of the cell, cell area, cell volume, and cell circularity are available for selection.</li>
		<li>Select the dataset to be displayed, such that every cell image can be visualized as a different data point on the scatter plot.</li>
		<li>Move the mouse cursor to each point, such that the corresponding image and other parameters are shown in a floating sub-window. 
		NOTE: The axes can be changed between the linear and logarithmic scales in an interactive manner.</li>
		<li>Perform further analysis of any subset of the dataset by manually gating on the scatter plot. 
		NOTE: The histogram in every parameter of the gated subset can be plotted against those of the entire library. The separate histograms are displayed in another floating sub-window.</li>
	</ol>
	</li>
</ol>

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<li>Wu, J., et al. <a target="_blank" href="https://www.eee.hku.hk/optima/pub/journal/1701_LSA.pdf">Ultrafast Laser-Scanning Time-Stretch Imaging at Visible Wavelengths.</a> Light Sci Appl. , (2016).</li>
</ol>

The authors have nothing to disclose.

There are several technical details that require special attention during the ATOM system setup. First, it is essential to note that asymmetric/oblique spectrally-encoded illumination could introduce residual phase-gradient components (i.e., the shadowing effect) in the absorption contrast and influence the enhancement of phase-gradient contrast in ATOM. Therefore, this effect of oblique illumination should be minimized. Second, it should be emphasized that the time-multiplexing or time-interleaving schemeinvolv...

Optical imaging presents a powerful tool and cell-based assay to (almost) non-invasively visualize the detailed spatial distribution of many cellular/subcellular components, thus uncovering a multitude of morphological, biophysical, and biomolecular signatures of cells. However, this ability to extract high-content information from single cells has generally been compromised when an enormous and heterogeneous population of cells had to be investigated. This marks a common trade-off in cell-based assays between measurement throughput and content. A notable example is that adding imaging capability to flow cytometry has resulted in a down-scaling of throughput by at least 1-2 orders of magnitude compared to that of the classical non-imaging flow cytometers. Although it could offer complex morphological single-cell analysis that is not possible with standard flow cytometers1, imaging flow cytometry generally lacks sufficient throughput to identify cellular heterogeneity with high statistical confidence. This is necessary for new discoveries in biology and for gaining an understanding of the pathogenesis of diseases. The key technical challenge lies in the inherent speed limit imposed by the common optical imaging strategies: laser-beam scanning, (e.g., by galvanometric mirrors), and/or image sensors (e.g., charge-coupled device (CCD) and complementary metal-oxide semiconductor (CMOS)). The laser scanning speed is intrinsically restricted by the mechanical inertia of the scanning mirrors, whereas the frame rate of CCD or CMOS is limited by the fundamental trade-off between imaging speed and sensitivity (i.e., increasing the frame rate leads to reduced signal detection sensitivity, and vice versa).
Based on an all-optical, ultrafast image-encoding mechanism, optical time-stretch imaging has been demonstrated as an attractive platform for high-throughput imaging flow cytometers, without need of the conventional image sensors or mechanical laser scanning2,3. Detailed descriptions of the working principle of time-stretch imaging can be found in references4,5,6,7. In brief, it consists of two interchangeable mapping steps: (i) spectral encoding (wavelength-space mapping), in which the spatial coordinates of the imaged specimen are mapped to different wavelengths across the spectrum of the light-pulsed beam8,9, and (ii) a dispersive Fourier transformation (wavelength-time mapping)9, in which the wavelength components of individual laser pulses are transformed (stretched) via group velocity dispersion (GVD) into temporal (wavelength-swept) waveforms (Figure 1). An important feature of time-stretch imaging is optical amplification, which plays a critical role in combatting the loss of sensitivity due to ultrafast photodetection and GVD loss, thus enhancing the image signal-to-noise ratio (SNR) without being contaminated by the photodetector noise9. Since each laser pulse encodes a line-scan of the imaged specimen, which is orthogonal to the unidirectional flow of the cells, an effective line-scan rate is determined by the laser repetition rate, which is typically beyond 10 MHz. This ultrafast operation enables blur-free, single-cell image capture at a throughput of 10,000-100,000 cells/s (i.e., 10-100 times higher than conventional imaging flow cytometry). As a result, time-stretch imaging could find unique applications in high-throughput, single-cell, image-based screening, especially when there is a need for identifying unknown heterogeneity or rare/aberrant cells within a sizable population (thousands to millions of cells), such as rare cancer cell screening10 or micro-algae classification11.
Time-stretch imaging predominantly relies on bright-field (BF) image capture, from which the image contrast is generated through light scattering and absorption from the cells2,3,9,10,11. Such label-free, single-cell imaging capabilities could bypass the detrimental effects associated with the fluorescent labels, such as cytotoxicity and photobleaching, and yet provide valuable information for single-cell analysis based on the cellular and sub-cellular texture and morphology. These label-free parameters are proven to be effective for the deep image classification of cells, especially when an enormous cell population is available11,12. However, in many occasions, BF imaging fails to provide sufficient contrast to reveal the detailed morphology of the label-free transparent cells. Different label-free, phase-contrast, time-stretch imaging modalities have been developed for enhancing the imaging contrast at ultrafast frame rates13,14,15. Among these techniques, asymmetric-detection time-stretch optical microscopy (ATOM) was developed to reveal the phase-gradient (differential-interference-contrast-(DIC)-like) contrast based on a concept similar to Schlieren photography, enabling the label-free, high-contrast imaging of single cells at an ultrahigh microfluidic speed (up to 10 m/s)16. This effect can be readily generated through oblique detection or illumination by partially blocking the image-encoded beam path or tilting the beam before photodetection. Another advantage of ATOM is its ability to simultaneously acquire two phase-gradient contrasts along opposite orientations. Intensity subtraction and summation of two opposite-contrast images yield the differential phase-gradient contrast and the absorption contrast, respectively, from the same line-scan. This work presents a detailed protocol describing the implementation of ATOM, including the establishment of the optical setup, the sample preparation, and the data acquisition and visualization. Specifically, this work demonstrates the ATOM operation with single-cell imaging of human blood cells, cancer cells, and phytoplankton (microalgae). This highlights the applicability of ATOM to imaging flow cytometry, not only in the biomedical arena, but also in marine and biofuel research17,18.

<table><tbody><tr><td>Nikon Plan Fluorite Physiology Objectives 40X</td><td>Nikon</td><td>MRF07420</td><td>Objective lens (Obj2)</td></tr><tr><td>Olympus Plan Fluorite Objective, 0.75 NA, 0.51 mm WD, 40X</td><td>Olympus</td><td>RMS40X-PF</td><td>Objective lens (Obj1)</td></tr><tr><td>N-BK7 Plano-Convex Lenses</td><td>Thorlabs</td><td>LA1145-C</td><td>For relaying spectral shower</td></tr><tr><td>FC/APC Fiber Collimation Package</td><td>Thorlabs</td><td>F220APC-1064</td><td>For outputing and collecting laser pulses</td></tr><tr><td>Pellicle beamsplitter</td><td>Thorlabs</td><td>BP145B3</td><td>For making beam replica</td></tr><tr><td>Protected silver mirror</td><td>Thorlabs</td><td>PF10-03-P01</td><td>For reflecting light</td></tr><tr><td>800-1650nm 12GHz single mode DC-coupled NIR Photoreceiver</td><td>Newport</td><td>1544-B</td><td>For converting light into electrical signal</td></tr><tr><td>Infiniium High-Performance Oscilloscope</td><td>Agilent</td><td>DSOX91604A</td><td>To save the light-converted electrical signals</td></tr><tr><td>HI1060 optical fiber</td><td>Corning</td><td>HI1060</td><td>Optical fiber for time-stretch</td></tr><tr><td>YTTERBIUM DOPED FIBER AMPLIFIER</td><td>Keopsys</td><td>KPS-STD-BT-YFA-37-BO-SM-111-FA-FA</td><td>Optical in-line amplifier</td></tr><tr><td>Holographic grating</td><td>Wasatch Photonics</td><td>020305-6</td><td>Grating</td></tr><tr><td>Infuse/Withdraw Syringe Pumps</td><td>Harvard Apparatus</td><td>PHD 2000</td><td>Syringe pump for sample loading in micro-fluidic channels</td></tr></tbody></table>

microfluidic imaging flow cytometry by asymmetric-detection time-stretch optical microscopy (atom)

Microfluidic Imaging Flow Cytometry by Asymmetric-detection Time-stretch Optical Microscopy (ATOM)

Scaling the number of measurable parameters, which allows for multidimensional data analysis and thus higher-confidence statistical results, has been the main trend in the advanced development of flow cytometry. Notably, adding high-resolution imaging capabilities allows for the complex morphological analysis of cellular/sub-cellular structures. This is not possible with standard flow cytometers. However, it is valuable for advancing our knowledge of cellular functions and can benefit life science research, clinical diagnostics, and environmental monitoring. Incorporating imaging capabilities into flow cytometry compromises the assay throughput, primarily due to the limitations on speed and sensitivity in the camera technologies. To overcome this speed or throughput challenge facing imaging flow cytometry while preserving the image quality, asymmetric-detection time-stretch optical microscopy (ATOM) has been demonstrated to enable high-contrast, single-cell imaging with sub-cellular resolution, at an imaging throughput as high as 100,000 cells/s. Based on the imaging concept of conventional time-stretch imaging, which relies on all-optical image encoding and retrieval through the use of ultrafast broadband laser pulses, ATOM further advances imaging performance by enhancing the image contrast of unlabeled/unstained cells. This is achieved by accessing the phase-gradient information of the cells, which is spectrally encoded into single-shot broadband pulses. Hence, ATOM is particularly advantageous in high-throughput measurements of single-cell morphology and texture &#8211; information indicative of cell types, states, and even functions. Ultimately, this could become a powerful imaging flow cytometry platform for the biophysical phenotyping of cells, complementing the current state-of-the-art biochemical-marker-based cellular assay. This work describes a protocol to establish the key modules of an ATOM system (from optical frontend to data processing and visualization backend), as well as the workflow of imaging flow cytometry based on ATOM, using human cells and micro-algae as the examples.

This work illustrates two single-cell imaging demonstrations by ATOM: one with mammalian cells (human peripheral blood mononuclear cells (PBMC) and breast cancer cells (MCF-7)) and another with phytoplanktons (Scenedesmus and Chlamydomonas). The first experiment was motivated by the growing interest in liquid biopsy for the detection, enumeration, and characterization of circulating tumor cells (CTCs) in the blood23. The ability to...

This protocol describes the implementation of an asymmetric-detection time-stretch optical microscopy system for single-cell imaging in ultrafast microfluidic flow and its applications in imaging flow cytometry.

Scaling the number of measurable parameters, which allows for multidimensional data analysis and thus higher-confidence statistical results, has been the ...

the overall goal of this protocol is to prepare and use an asymmetric detection time stretch optical microscopy system for label-free high contrast single-cell imaging in ultra-fast microfluidic flow. The main advantage of ATOM is that it's allows ultra-fast image capture with line scan rates of up to tens of megahertz while enhancing the image contrast on the unlabeled cells. ATOM enables complex morphological analysis of cellular and subcellular structures with an uncentered cell-based assays at up to 100, 000 cells per second, which is not possible with standard flow cytometry.to begin setting up the system use a fiber collimator to couple a broadband femto or pico second near IR pulsed laser to a single mode dispersive optical fiber connected to an optical amplifier. Illuminate a transmission diffraction grading with the laser to produce a spectral shower. Orient the grading close to the littrow configuration to maximize the diffraction efficiency.Configure a telescopic relay lens assembly in a 4f imaging system to direct the spectral shower onto the rear focal plane of an objective lens below a sample platform. Ensure that the spectral shower fills the rear aperture of the objective lens. Place another objective lens with a plane mirror at its rear aperture above the sample platform.Adjust the objective lenses until their focal planes overlap. Verify that the spectral shower double passes the image plane and follows the same path back to the grading. Place a microfluidic chip on the sample platform and ensure that the spectral shower falls across the microfluidic channel.Redirect the returned light with a beam splitter after it passes through the grading. Separate the returned light into two beams with another beam splitter. Use fiber collimators to couple the beams to single mode fibers of different lengths to introduce a time delay between beams.Connect the fibers to a real-time oscilloscope through a fiber coupler and a photodetector. Tune the optical amplifier gain until the signal-to-noise ratio of the optical signal is greater than 10 decibels. Then block approximately half of each returned beam with knife-edge beam blocks.Adjust the beam blocks until the measured optical power in each fiber is about half of the power before the block. Verify that the oscilloscope bandwidth and sampling rate are both high enough not to influence the image resolution. To begin the experiment dilute cell samples to between 10 to the 5th and 10 to the 6th cells per milliliter with phosphate buffered saline or spring water.Then load the cell solution into a 10 milliliter syringe. Connect the syringe to the microfluidic channel inlet. Direct the microfluidic channel outlet to a centrifuge tube.Secure the syringe in a syringe pump and set the flow rate appropriately for the channel width. Choose the number of data points to be saved for the experiment. Then acquire and save the optical signal data.Digitize and export the data from the oscilloscope to a computer. Process the data into 2D images and extract a library of parameters from the images. Import the images in parameter library into a visualization program.Assign a parameter of interest to each axis and select the data set to be displayed as a scatter plot. Check the cell images associated with each group in the plot to identify trends. Use manual gating or display additional datasets for further analysis as needed.ATOM was used to acquire high contrast single-cell images of human cells and phytoplankton. Characteristic cell structures such as vacuoles, pyrenoids, and flagella, are visible in the phytoplankton images. Parameters such as cell size and circularity were determined from the ATOM images to classify cells for cytometric analysis.The classification results can be displayed as 2D scatter plots. Here volume versus circularity was plotted for an MCF7 shown in yellow and PBMC shown in red. Two clusters were observed for MCF7.By identifying the high contrast images corresponding to individual data points, the lower cluster was found to correspond to cell fragments and debris. Once mastered the imaging system can be set up in two to three hours if it is performed properly. The imaging procedures can be done within an hour.Don't forget to pay special attention to eye safety while working with the laser source and laser beam alignment. Precautions such as wearing safety goggles should always be taken while performing these procedures. While attempting this procedure remember to monitor the optical beam alignment throughout the whole imaging system setup process to ensure optimal imaging quality.While this technique can lend insight into the detection and quantification of rare or apparent cells during disease process, it can also be applied to cell type classification, within large and heterogeneous cell populations. This technique is particularly advantageous when looking to overcome the throughput limitation of imaging cytometry, while preserving the image quality. After watching this video, you should have good understanding of how to implement ATOM from the system setup to the imaging procedures.

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Engineering

In this article we describe how we obtain FRET traces from individual DNA molecules immobilized to a surface using an automated scanning confocal microscope.

Automated System for Single Molecule Fluorescence Measurements of Surface-immobilized Biomolecules

Droplet-based microfluidic platforms are promising candidates for high throughput experimentation since they are able to generate picoliter, self-compartmentalized vessels inexpensively at kHz rates. Through integration with fast, sensitive and high resolution fluorescence spectroscopic methods, the large amounts of information generated within these systems can be efficiently extracted, harnessed and utilized.

Fluorescence detection methods for microfluidic droplet platforms

We review our recent results on the integration of fluorescent microscopy and imaging flow cytometry tools on a cell-phone using compact and cost-effective opto-fluidic attachments. These cell-phone based micro-analysis devices might be useful for cytometric analysis, such as performing various cell counting tasks as well as for high-throughput screening of e.g., water samples in resource limited settings.

Wide-field Fluorescent Microscopy and Fluorescent Imaging Flow Cytometry on a Cell-phone

Confocal microscopy is used to image quiescent and flowing colloid-polymer mixtures, which are studied as model systems for attractive suspensions. Image analysis algorithms are used to calculate structural and dynamic metrics for the colloidal particles that measure changes due to geometric confinement.

Confocal Imaging of Confined Quiescent and Flowing Colloid-polymer Mixtures

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.

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

The in vitro micronucleus assay is a well-established method for evaluating genotoxicity and cytotoxicity but scoring the assay using manual microscopy is laborious and suffers from subjectivity and inter-scorer variability. This paper describes the protocol developed to perform a fully automated version of the assay using multispectral imaging flow cytometry.

An Automated Method to Perform The In Vitro Micronucleus Assay using Multispectral Imaging Flow Cytometry

We present protocols for simple actin filament microfluidic assays, in combination with fluorescence microscopy, that allow one to accurately monitor individual actin filaments in real-time while sequentially exposing them to different protein solutions.

Using Microfluidics and Fluorescence Microscopy to Study the Assembly Dynamics of Single Actin Filaments and Bundles

Microfluidics is a powerful tool for the development of diagnostic tests. However, expensive equipment and materials, as well as laborious fabrication and handling techniques, are often required. Here, we detail the fabrication protocol of an acrylic microfluidic device for magnetic micro- and nanoparticle-based immunoassays in a low-cost and simple-to-use setting.

Computer Numerical Control Micromilling of a Microfluidic Acrylic Device with a Staggered Restriction for Magnetic Nanoparticle-Based Immunoassays

Visual, single-molecule biochemistry studied through microfluidic chambers is greatly facilitated using glass barrel, gas-tight syringes, stable connections of tubing to flow cells, and elimination of bubbles by placing switching valves between the syringes and tubing. The protocol describes dual optical traps that enable visualization of DNA transactions and intermolecular interactions.

Use of Dual Optical Tweezers and Microfluidics for Single-Molecule Studies

This protocol describes a quantitative approach to measure microbial autoaggregation using imaging flow cytometry.

Imaging Flow Cytometry to Study Microbial Autoaggregation

The protocol describes an advanced microfluidic platform to quantitatively measure cytokine secretion dynamics of individual human peripheral blood mononuclear cells. The platform measures up to three cytokines in parallel (IL-6, TNFα, and IL-1β) for each individual cell stimulated with lipopolysaccharide as an example.

Microfluidic Approach to Resolve Simultaneous and Sequential Cytokine Secretion of Individual Polyfunctional Cells

Learn to implement ATOM for label-free high contrast single-cell imaging in ultrafast microfluidic flow. This method enables imaging at 100,000 cells per second with enhanced contrast for unlabeled cells.

Microfluidic Imaging Flow Cytometry by Asymmetric-detection Time-stretch Optical Microscopy (ATOM)

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