A. Ozcan gratefully acknowledges the support of the Presidential Early Career Award for Scientists and Engineers (PECASE), Army Research Office (ARO) Young Investigator Award, National Science Foundation (NSF) CAREER Award, Office of Naval Research (ONR) Young Investigator Award and National Institutes of Health (NIH) Director's New Innovator Award DP2OD006427 from the Office of the Director, National Institutes of Health.

In this section, we introduce the experimental protocols for our cell-phone based wide-field fluorescence microscopy13 and opto-fluidic imaging cytometry platform14. We will use fluorescent beads and fluorescently labeled white blood cells to test these imaging platforms.
A. Preparation of Cell-phone Based Wide-field Fluorescent Microscope and Opto-fluidic Imaging Flow Cytometer
The cell-phone based wide-field fluorescent microscope or flow cytometer consists of two major parts: a camera phone and a compact opto-fluidic add-on attachment.
1. The Camera Phone
While the presented techniques are applicable to almost any camera phone, we have chosen Sony Erickson Aino as the base for these devices. This cell-phone has an ~ 8 MegaPixel RGB CMOS sensor installed on it and a built-in lens that has a focal length (f1) of ~ 4.65 mm.
2. Opto-fluidic Attachment for Wide-field Fluorescent Microscopy
The optical attachment is designed by Autodesk and is printed by a Dimension Elite 3-D printer using ABSplus thermoplastic material. In this printing process, model and support materials are heated in an extrusion head within the printer and are deposited layer by layer on a modeling base. When this step is completed, the supporting material can be dissolved, leaving a robust 3D model of the desired prototype Our optical attachment design consists of LEDs (center wavelength at ~ 470 nm, Digikey), a plastic filter (#NT54-46, Edmund Optics), a sample tray, and a plano-convex lens f2=15 mm (# NT45-302, Edmund Optics). All the LEDs and plastic filters can be easily changed based on the fluorophores' spectra. The steps for assembling this opto-fluidic attachment include:
<ol>
	<li>Place the lens into the attachment within its specific lens holder position.</li>
	<li>Place the plastic filter onto the filter tray and slide it into the attachment; or tape the plastic filter in front of the cell-phone camera lens.</li>
	<li>Insert the LED tray into the attachment.</li>
	<li>Place the sample glass slides into the sample tray. Slide the sample tray into the attachment. Face the LEDs toward the sample.</li>
	<li>Clip the attachment onto the cell-phone, such that the extra lens is directly in touch with the cell-phone camera lens.</li>
	<li>Use the switch on the attachment to turn on the LEDs.</li>
	<li>Image the sample of interest with the cell-phone camera unit using its "night mode".</li>
</ol>
3. Opto-fluidic Attachment for Fluorescent Imaging Cytometry
When there is a need to screen large volumes of liquid samples for the detection of rare events, optofluidic flow cytometry device could be preferred. We can modify our wide-field fluorescent microscope design and convert it into a flow cytometer, where a PDMS based microfluidic channel is used to continually deliver the liquid sample through the imaging volume. The optical attachment is also designed by Autodesk and printed by Dimension Elite 3-D printer. It also consists of LEDs (center wavelength at ~ 470 nm, Digikey), a plastic filter (#NT54-46, Edmund Optics), a sample tray, and an aspherical lens (f = 4.5 mm) (product # C230TME-A; Thorlab). The steps for assembling this opto-fluidic attachment include:
<ol>
	<li>Place the aspherical lens into the attachment.</li>
	<li>Place the plastic filter onto the filter tray and slide it into the attachment; or tape the plastic filter in front of the cellphone camera lens.</li>
	<li>Slide the microfluidic channel into the same opto-fluidic attachment.</li>
	<li>Clip the attachment onto the cell-phone such that the extra lens is directly in touch with the cell-phone camera lens.</li>
	<li>Use the switch on the attachment to turn on the LEDs.</li>
	<li>Connect the microfluidic channel to the syringe pump and deliver the liquid sample into the microfluidic device at a constant flow rate.</li>
	<li>Capture a movie of the fluorescent cells/particles flowing through the microfluidic channel using the video mode of the cell-phone camera.</li>
</ol>
B. Sample Preparation
4. Preparation of Fluorescent Micro-particle Samples
<ol>
	<li>Fluorescent beads with 10 &mu;m diameter (red beads: product #F8834 excitation/emission 580nm/605nm; green beads: product #F8836: excitation/emission 505nm/515nm) are purchased from Invitrogen (Carlsbad, CA).</li>
	<li>Mix 10 &mu;l of green fluorescent beads, 10 &mu;l of red fluorescent beads with 40 &mu;l of DI water.</li>
	<li>Place10 &mu;l of this bead mixture on a glass slide using a micropipette and put another glass slide on the top of it to make a sandwich structure.</li>
	<li>Insert this sandwich structure into the sample tray and slide it into the cell-phone attachment.</li>
</ol>
5. Preparation of Fluorescently Labeled White Blood Cells
<ol>
	<li>Take SYTO16 nucleic acid fluorescent labeling kit (#S7578, Life Technology) and phosphate buffered saline (PBS) out from the refrigerator and bring them to room temperature.</li>
	<li>Transfer 200 &mu;l whole blood sample from EDTA blood collection tube to 1.5 ml polystyrene tube (# 05-408-129, Fisher Scientific).</li>
	<li>Add 1 ml red blood cell lysing buffer (# R7757, Sigma-Aldrich) to the 200 &mu;l whole blood sample and mix thoroughly.</li>
	<li>After 5 min, centrifuge the lysed blood sample and remove the supernatant solution.</li>
	<li>Resuspend the white blood cell pellet into 200 &mu;l PBS buffer and gently mix them.</li>
	<li>Add 5 &mu;l 1 mM SYTO16 solution to the white blood cell sample. Wrap the sample with aluminum foil and incubate in dark environment for ~ 30 min.</li>
	<li>Centrifuge the sample again. Supernatant is removed and the labeled white blood cell pellet is re-suspended into PBS buffer.</li>
	<li>Place 5-10 &mu;l labeled white blood cell liquid sample to a cover slip, and place a second cover slip on the top of the sample.</li>
	<li>Insert the sandwiched sample slide into the sample tray and image it using the cell-phone fluorescent microscope.</li>
</ol>
Alternatively
<ol start="10">
	<li>Continually deliver the fluorescently labeled white blood cells through a microfluidic channel using an automated syringe pump, while also capturing a fluorescent microscopic movie of the flowing cells using the cell-phone camera in video mode. We should also emphasize that a portable battery powered syringe pump or even gravity force can be utilized to drive the flow through the microfluidic channel.</li>
</ol>

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W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Toward+fluorescence+nanoscopy.">Toward fluorescence nanoscopy.</a> Nature Biotechnology. 21, 1347-1355 (2003).</li><li>Gustafsson, M. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nonlinear+structured-illumination+microscopy:+wide-field+fluorescence+imaging+with+theoretically+unlimited+resolution.">Nonlinear structured-illumination microscopy: wide-field fluorescence imaging with theoretically unlimited resolution.</a> Proceedings of the National Academy of Science U.S.A. 102, 13081-13086 (2005).</li><li>Betzig, E., Patterson, G. H., Sougrat, R., Lindwasser, O. W., Olenych, S., Bonifacino, J. S., Davidson, M. W., Lippincott-Schwartz, J., Hess, H. 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Dr. Ozcan is the founder of a start-up company that aims to commercialize computational imaging and microscopy tools.

We have presented our recent results on cell-phone based wide-field fluorescent microscopy and opto-fluidic imaging flow cytometry using light-weight and compact opto-fluidic attachments to cell-phone cameras. Using this platform technology we imaged fluorescent objects including micro-particles and labeled white blood cells in whole blood samples. Therefore, this compact and cost-effective cell-phone based fluorescent imaging toolset might be useful for point-of-care diagnosis, especially in resource limited regions for...

Microscopy and flow-cytometry are widely used techniques1-12 in biomedical and scientific research as well as clinical diagnosis for counting and characterization of various cell types. However, conventional microscopes and flow-cytometry instruments are relatively complex and expensive, which limits their use to mainly well-established central laboratories. Recently we have developed a compact and lightweight fluorescent imaging cytometry and microscopy device integrated on a cell-phone,13,14 which shows promise to cost-effectively translate fluorescent microscopy, flow-cytometry and related micro-analysis techniques to resource-limited environments for various telemedicine applications impacting global health.
In the optofluidic flow-cytometry configuration (see e.g. Figure 1C and 1D), a custom-designed polydimethysiloxane (PDMS) based microfluidic channel is positioned in front of the cell-phone camera-unit, where light-emitting-diodes (LEDs) are butt-coupled to the edges of the channel. This microfluidic chip, together with the liquid sample inside, forms an opto-fluidic planar waveguide (composed of for example PDMS-liquid-PDMS) such that the excitation light is guided to uniformly pump the fluorescent labeled specimens inside the micro-channel. The fluorescence emission from these labeled objects, e.g. cells, is further imaged through an additional lens placed right after the cell-phone camera-unit and is mapped onto the cell-phone Complementary Metal-Oxide-Semiconductor (CMOS) image sensor. Since the fluorescent emission is collected perpendicular to the excitation light path, an inexpensive plastic absorption filter is sufficient to remove the scattered excitation light and can provide a decent dark-field background required for fluorescent imaging. Using a similar opto-fluidic design, we can also image the fluorescent objects in static mode (see Figure 1A and 1B), in which the fluorescent particles are sandwiched between two glass slides instead of flowing through a microfluidic channel and the fluorescent emission from these fluorescent particles are captured by the cell-phone CMOS image sensor for particle counting and characterization. Based on different application requirements, flow cytometry or wide-field fluorescent microscopy can be chosen. For example, cell-phone flow cytometry device could be especially useful for screening large volumes of liquid samples (e.g. a few ml) for the detection of rare cells or pathogens.
In this manuscript we review some of 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, imaging cytometry and sensing platforms could provide various opportunities for telemedicine and point-of-care diagnostics, especially impacting our fight against global health challenges in resource limited regions of the world.

<table border="1">
	<tbody>
		<tr>
			<td>Name </td>
			<td>Company</td>
			<td>Catalog Number </td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Cell-phone</td>
			<td>Sony</td>
			<td>Sony Ericsson Aino</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Plano-convex lens</td>
			<td>Edmund Optics</td>
			<td># NT45-302</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Aspherical lens</td>
			<td>Thorlab</td>
			<td># C230TME-A</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Filter</td>
			<td>Edmund Optics</td>
			<td>#NT54-46</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Blue LED</td>
			<td>Digikey</td>
			<td>#365-1201-ND</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Battery</td>
			<td>Digikey</td>
			<td>#P032-ND</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Polystyrene tube</td>
			<td>Fisher Scientific</td>
			<td>#05-408-129</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Red blood cell lysing buffer</td>
			<td>Sigma Aldrich</td>
			<td>R7757</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>SYLGARD 184 SILICONE ELASTOMER KIT</td>
			<td>Dow Corning</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Red fluorescent beads (10 &mu;m)</td>
			<td>Life Technologies</td>
			<td>#F8834</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Green fluorescent beads (10 &mu;m)</td>
			<td>Life Technologies</td>
			<td>#F8836</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>SYTO16 nucleic acid fluorescent labeling</td>
			<td>Life Technologies</td>
			<td># S7578</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>

wide-field fluorescent microscopy and fluorescent imaging flow cytometry on a cell-phone

Fluorescent microscopy and flow cytometry are widely used tools in biomedical research and clinical diagnosis. However these devices are in general relatively bulky and costly, making them less effective in the resource limited settings. To potentially address these limitations, we have recently demonstrated the integration of wide-field fluorescent microscopy and imaging flow cytometry tools on cell-phones using compact, light-weight, and cost-effective opto-fluidic attachments. In our flow cytometry design, fluorescently labeled cells are flushed through a microfluidic channel that is positioned above the existing cell-phone camera unit. Battery powered light-emitting diodes (LEDs) are butt-coupled to the side of this microfluidic chip, which effectively acts as a multi-mode slab waveguide, where the excitation light is guided to uniformly excite the fluorescent targets. The cell-phone camera records a time lapse movie of the fluorescent cells flowing through the microfluidic channel, where the digital frames of this movie are processed to count the number of the labeled cells within the target solution of interest. Using a similar opto-fluidic design, we can also image these fluorescently labeled cells in static mode by e.g. sandwiching the fluorescent particles between two glass slides and capturing their fluorescent images using the cell-phone camera, which can achieve a spatial resolution of e.g. ~ 10 &mu;m over a very large field-of-view of ~ 81 mm2. This cell-phone based fluorescent imaging flow cytometry and microscopy platform might be useful especially in resource limited settings, for e.g. counting of CD4+ T cells toward monitoring of HIV+ patients or for detection of water-borne parasites in drinking water.

With our opto-fluidic pumping/excitation scheme (Figures 1C and 1D), fluorescently labeled cells can be continuously delivered into the microfluidic channel using a syringe pump while the cell-phone camera records a time-lapse fluorescent microscopic movie of the flowing cells. These fluorescent movies can then be rapidly analyzed using contour-detection and tracking algorithms14,15 to automatically determine the absolute number and the density of the cells flowing through the...

Watch this Scientific Journal Video about Wide-field Fluorescent Microscopy and Fluorescent Imaging Flow Cytometry on a Cell-phone at JoVE.com

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

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.

Fluorescent microscopy and flow cytometry are widely used tools in biomedical research and clinical diagnosis. However these devices are in general ...

wide-field-fluorescent-microscopy-fluorescent-imaging-flow-cytometry

Research

JoVE Journal

Bioengineering

23.5K ビュー.  University of California, Los Angeles. 我々は、コンパクトでコスト効果の高い光流体添付ファイルを使用して携帯電話上の蛍光顕微鏡やイメージングフローサイトメトリーツールの統合、我々の最近の結果を確認します。これらの携帯電話基にしたマイクロ分析装置は、以下のようなリソースが限られた設定で、例えば、水試料のハイスループットスクリーニングのためだけでなく、様々な?...

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

Fluorescent Nanoparticles for the Measurement of Ion Concentration in Biological Systems

Tightly regulated ion homeostasis throughout the body is necessary for the prevention of such debilitating states as dehydration.1 In contrast, rapid ion fluxes at the cellular level are required for initiating action potentials in excitable cells.2 Sodium regulation plays an important role in both of these cases; however, no method currently exists for continuously monitoring sodium levels in vivo 3 and intracellular sodium probes 4 do not provide similar detailed results as calcium probes. In an effort to fill both of these voids, fluorescent nanosensors have been developed that can monitor sodium concentrations in vitro and in vivo.5,6 These sensors are based on ion-selective optode technology and consist of plasticized polymeric particles in which sodium specific recognition elements, pH-sensitive fluorophores, and additives are embedded.7-9 Mechanistically, the sodium recognition element extracts sodium into the sensor. 10 This extraction causes the pH-sensitive fluorophore to release a hydrogen ion to maintain charge neutrality within the sensor which causes a change in fluorescence. The sodium sensors are reversible and selective for sodium over potassium even at high intracellular concentrations.6 They are approximately 120 nm in diameter and are coated with polyethylene glycol to impart biocompatibility. Using microinjection techniques, the sensors can be delivered into the cytoplasm of cells where they have been shown to monitor the temporal and spatial sodium dynamics of beating cardiac myocytes.11 Additionally, they have also tracked real-time changes in sodium concentrations in vivo when injected subcutaneously into mice.3 Herein, we explain in detail and demonstrate the methodology for fabricating fluorescent sodium nanosensors and briefly demonstrate the biological applications our lab uses the nanosensors for: the microinjection of the sensors into cells; and the subcutaneous injection of the sensors into mice.

Fluorescent nanoparticles produced in our lab are used for imaging ion concentrations and ion fluxes in biological systems such as cells during signaling and interstitial fluid during physiological homeostasis.

Tightly regulated ion homeostasis throughout the body is necessary for the prevention of such debilitating states as dehydration.1 In contrast, rapid ion ...

Lensless Fluorescent Microscopy on a Chip

On-chip lensless imaging in general aims to replace bulky lens-based optical microscopes with simpler and more compact designs, especially for high-throughput screening applications. This emerging technology platform has the potential to eliminate the need for bulky and/or costly optical components through the help of novel theories and digital reconstruction algorithms. Along the same lines, here we demonstrate an on-chip fluorescent microscopy modality that can achieve e.g., &lt;4&mu;m spatial resolution over an ultra-wide field-of-view (FOV) of &gt;0.6-8 cm2 without the use of any lenses, mechanical-scanning or thin-film based interference filters. In this technique, fluorescent excitation is achieved through a prism or hemispherical-glass interface illuminated by an incoherent source. After interacting with the entire object volume, this excitation light is rejected by total-internal-reflection (TIR) process that is occurring at the bottom of the sample micro-fluidic chip. The fluorescent emission from the excited objects is then collected by a fiber-optic faceplate or a taper and is delivered to an optoelectronic sensor array such as a charge-coupled-device (CCD). By using a compressive-sampling based decoding algorithm, the acquired lensfree raw fluorescent images of the sample can be rapidly processed to yield e.g., &lt;4&mu;m resolution over an FOV of &gt;0.6-8 cm2. Moreover, vertically stacked micro-channels that are separated by e.g., 50-100 &mu;m can also be successfully imaged using the same lensfree on-chip microscopy platform, which further increases the overall throughput of this modality. This compact on-chip fluorescent imaging platform, with a rapid compressive decoder behind it, could be rather valuable for high-throughput cytometry, rare-cell research and microarray-analysis.

A lensless on-chip fluorescent microscopy platform is demonstrated that can image fluorescent objects over an ultra-wide field-of-view of e.g., &#62;0.6-8 cm2 with &#60;4&#956;m resolution using a compressive sampling based decoding algorithm. Such a compact and wide-field fluorescent on-chip imaging modality could be valuable for high-throughput cytometry, rare-cell research and microarray-analysis.

On-chip lensless imaging in general aims to replace bulky lens-based optical microscopes with simpler and more compact designs, especially for ...

Specimen Preparation, Imaging, and Analysis Protocols for Knife-edge Scanning Microscopy

Major advances in high-throughput, high-resolution, 3D microscopy techniques have enabled the acquisition of large volumes of neuroanatomical data at submicrometer resolution. One of the first such instruments producing whole-brain-scale data is the Knife-Edge Scanning Microscope (KESM)7, 5, 9, developed and hosted in the authors' lab. KESM has been used to section and image whole mouse brains at submicrometer resolution, revealing the intricate details of the neuronal networks (Golgi)1, 4, 8, vascular networks (India ink)1, 4, and cell body distribution (Nissl)3. The use of KESM is not restricted to the mouse nor the brain. We have successfully imaged the octopus brain6, mouse lung, and rat brain. We are currently working on whole zebra fish embryos. Data like these can greatly contribute to connectomics research10; to microcirculation and hemodynamic research; and to stereology research by providing an exact ground-truth. 
In this article, we will describe the pipeline, including specimen preparation (fixing, staining, and embedding), KESM configuration and setup, sectioning and imaging with the KESM, image processing, data preparation, and data visualization and analysis. The emphasis will be on specimen preparation and visualization/analysis of obtained KESM data. We expect the detailed protocol presented in this article to help broaden the access to KESM and increase its utilization.

The full process from brain specimen preparation to serial sectioning imaging using the Knife-Edge Scanning Microscope, to data visualization and analysis is described. This technique is currently used to acquire mouse brain data, but it is applicable to other organs, other species.

Major advances in high-throughput, high-resolution, 3D microscopy techniques have enabled the acquisition of large volumes of neuroanatomical data at ...

Analyzing Cellular Internalization of Nanoparticles and Bacteria by Multi-spectral Imaging Flow Cytometry

Nanoparticulate systems have emerged as valuable tools in vaccine delivery through their ability to efficiently deliver cargo, including proteins, to antigen presenting cells1-5. Internalization of nanoparticles (NP) by antigen presenting cells is a critical step in generating an effective immune response to the encapsulated antigen. To determine how changes in nanoparticle formulation impact function, we sought to develop a high throughput, quantitative experimental protocol that was compatible with detecting internalized nanoparticles as well as bacteria. To date, two independent techniques, microscopy and flow cytometry, have been the methods used to study the phagocytosis of nanoparticles. The high throughput nature of flow cytometry generates robust statistical data. However, due to low resolution, it fails to accurately quantify internalized versus cell bound nanoparticles. Microscopy generates images with high spatial resolution; however, it is time consuming and involves small sample sizes6-8. Multi-spectral imaging flow cytometry (MIFC) is a new technology that incorporates aspects of both microscopy and flow cytometry that performs multi-color spectral fluorescence and bright field imaging simultaneously through a laminar core. This capability provides an accurate analysis of fluorescent signal intensities and spatial relationships between different structures and cellular features at high speed.
	Herein, we describe a method utilizing MIFC to characterize the cell populations that have internalized polyanhydride nanoparticles or Salmonella enterica serovar Typhimurium. We also describe the preparation of nanoparticle suspensions, cell labeling, acquisition on an ImageStreamX system and analysis of the data using the IDEAS application. We also demonstrate the application of a technique that can be used to differentiate the internalization pathways for nanoparticles and bacteria by using cytochalasin-D as an inhibitor of actin-mediated phagocytosis.

In this article, we describe a method utilizing multi-spectral imaging flow cytometry to quantify the internalization of polyanhydride nanoparticles or bacteria by RAW 264.7 cells.

Nanoparticulate  systems have emerged as valuable tools in vaccine delivery through their  ability to efficiently deliver cargo, including proteins, to ...

Localization and Relative Quantification of Carbon Nanotubes in Cells with Multispectral Imaging Flow Cytometry

Carbon-based nanomaterials, like carbon nanotubes (CNTs), belong to this type of nanoparticles which are very difficult to discriminate from carbon-rich cell structures and de facto there is still no quantitative method to assess their distribution at cell and tissue levels. What we propose here is an innovative method allowing the detection and quantification of CNTs in cells using a multispectral imaging flow cytometer (ImageStream, Amnis). This newly developed device integrates both a high-throughput of cells and high resolution imaging, providing thus images for each cell directly in flow and therefore statistically relevant image analysis. Each cell image is acquired on bright-field (BF), dark-field (DF), and fluorescent channels, giving access respectively to the level and the distribution of light absorption, light scattered and fluorescence for each cell. The analysis consists then in a pixel-by-pixel comparison of each image, of the 7,000-10,000 cells acquired for each condition of the experiment. Localization and quantification of CNTs is made possible thanks to some particular intrinsic properties of CNTs: strong light absorbance and scattering; indeed CNTs appear as strongly absorbed dark spots on BF and bright spots on DF with a precise colocalization.
This methodology could have a considerable impact on studies about interactions between nanomaterials and cells given that this protocol is applicable for a large range of nanomaterials, insofar as they are capable of absorbing (and/or scattering) strongly enough the light.

In this study, the main purpose was to monitor the cellular uptake and eventual exocytosis of carbon nanotubes (CNTs). From this perspective, we proposed here a unique method using multispectral imaging flow cytometry allowing a quantification and localization of CNTs in a statistically relevant number of cells.

Carbon-based nanomaterials, like carbon nanotubes (CNTs), belong to this type of nanoparticles which are very difficult to discriminate from carbon-rich ...

A Novel Method for Localizing Reporter Fluorescent Beads Near the Cell Culture Surface for Traction Force Microscopy

PA gels have long been used as a platform to study cell traction forces due to ease of fabrication and the ability to tune their elastic properties. When the substrate is coated with an extracellular matrix protein, cells adhere to the gel and apply forces, causing the gel to deform. The deformation depends on the cell traction and the elastic properties of the gel. If the deformation field of the surface is known, surface traction can be calculated using elasticity theory. Gel deformation is commonly measured by embedding fluorescent marker beads uniformly into the gel. The probes displace as the gel deforms. The probes near the surface of the gel are tracked. The displacements reported by these probes are considered as surface displacements. Their depths from the surface are ignored. This assumption introduces error in traction force evaluations. For precise measurement of cell forces, it is critical for the location of the beads to be known. We have developed a technique that utilizes simple chemistry to confine fluorescent marker beads, 0.1 and 1 &#181;m in diameter, in PA gels, within 1.6 &#956;m of the surface. We coat a coverslip with poly-D-lysine (PDL) and fluorescent beads. PA gel solution is then sandwiched between the coverslip and an adherent surface. The fluorescent beads transfer to the gel solution during curing. After polymerization, the PA gel contains fluorescent beads on a plane close to the gel surface.

Traditional techniques for fabricating polyacrylamide (PA) gels containing fluorescent probes involve sandwiching a gel between an adherent surface and a glass slide. Here, we show that coating this slide with poly-D-lysine (PDL) and fluorescent probes localizes the probes to within 1.6 &#181;m from the gel surface.

PA gels have long been used as a platform to study cell traction forces due to ease of fabrication and the ability to tune their elastic properties. When ...

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous number of imaging techniques currently available and the progress in instrumentation, there is still a need for highly sensitive probes that are suitable for in vivo imaging. One typical problem of available preclinical fluorescent probes is their rapid clearance in vivo, which reduces their imaging sensitivity. To circumvent rapid clearance, increase number of dye molecules at the target site, and thereby reduce background autofluorescence, encapsulation of the near-infrared fluorescent dye, DY-676-COOH in liposomes and verification of its potential for in vivo imaging of inflammation was done. DY-676 is known for its ability to self-quench at high concentrations. We first determined the concentration suitable for self-quenching, and then encapsulated this quenching concentration into the aqueous interior of PEGylated liposomes. To substantiate the quenching and activation potential of the liposomes we use a harsh freezing method which leads to damage of liposomal membranes without affecting the encapsulated dye. The liposomes characterized by a high level of fluorescence quenching were termed Lip-Q. We show by experiments with different cell lines that uptake of Lip-Q is predominantly by phagocytosis which in turn enabled the characterization of its potential as a tool for in vivo imaging of inflammation in mice models. Furthermore, we use a zymosan-induced edema model in mice to substantiate the potential of Lip-Q in optical imaging of inflammation in vivo. Considering possible uptake due to inflammation-induced enhanced permeability and retention (EPR) effect, an always-on liposome formulation with low, non-quenched concentration of DY-676-COOH (termed Lip-dQ) and the free DY-676-COOH were compared with Lip-Q in animal trials.

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

The use of fluorophores for in vivo imaging can be greatly limited by opsonization, rapid clearance, low detection sensitivity and cytotoxic effects on the host. Encapsulation of fluorophores in liposomes by film hydration and extrusion leads to fluorescence quenching and protection which enables in vivo imaging with high detection sensitivity.

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous ...

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

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

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

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

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

The in vitro micronucleus (MN) assay is often used to evaluate cytotoxicity and genotoxicity but scoring the assay via manual microscopy is laborious and introduces uncertainty in results due to variability between scorers. To remedy this, automated slide-scanning microscopy as well as conventional flow cytometry methods have been introduced in an attempt to remove scorer bias and improve throughput. However, these methods have their own inherent limitations such as inability to visualize the cytoplasm of the cell and the lack of visual MN verification or image data storage with flow cytometry. Multispectral Imaging Flow Cytometry (MIFC) has the potential to overcome these limitations. MIFC combines the high resolution fluorescent imagery of microscopy with the statistical robustness and speed of conventional flow cytometry. In addition, all collected imagery can be stored in dose-specific files. This paper describes the protocol developed to perform a fully automated version of the MN assay on MIFC. Human lymphoblastoid TK6 cells were enlarged using a hypotonic solution (75 mM KCl), fixed with 4% formalin and the nuclear content was stained with Hoechst 33342. All samples were run in suspension on the MIFC, permitting acquisition of high resolution images of all key events required for the assay (e.g. binucleated cells with and without MN as well as mononucleated and polynucleated cells). Images were automatically identified, categorized and enumerated in the MIFC data analysis software, allowing for automated scoring of both cytotoxicity and genotoxicity. Results demonstrate that using MIFC to perform the in vitro MN assay allows statistically significant increases in MN frequency to be detected at several different levels of cytotoxicity when compared to solvent controls following exposure of TK6 cells to Mitomycin C and Colchicine, and that no significant increases in MN frequency are observed following exposure to Mannitol.

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.

The in vitro micronucleus (MN) assay is often used to evaluate cytotoxicity and genotoxicity but scoring the assay via manual microscopy is laborious and ...

Ovarian Cancer Detection Using Photoacoustic Flow Cytometry

Many studies suggest that the enumeration of circulating tumor cells (CTCs) may show promise as a prognostic tool for ovarian cancer. Current strategies for the detection of CTCs include flow cytometry, microfluidic devices, and real-time polymerase chain reaction (RT-PCR). Despite recent advances, methods for the detection of early ovarian cancer metastasis still lack the sensitivity and specificity required for clinical translation. Here, a novel method is presented for the detection of ovarian circulating tumor cells by photoacoustic flow cytometry (PAFC) utilizing a custom three dimensional (3D) printed system, including a flow chamber and syringe pump. This method utilizes folic acid-capped copper sulfide nanoparticles (FA-CuS NPs) to target SKOV-3 ovarian cancer cells by PAFC. This work demonstrates the affinity of these contrast agents for ovarian cancer cells. The results show NP characterization, PAFC detection, and NP uptake by fluorescence microscopy, thus demonstrating the potential of this novel system to detect ovarian CTCs at physiologically relevant concentrations.

A protocol is presented to detect circulating ovarian tumor cells utilizing a custom-made photoacoustic flow system and targeted folic acid-capped copper sulfide nanoparticles.

Many studies suggest that the enumeration of circulating tumor cells (CTCs) may show promise as a prognostic tool for ovarian cancer. Current strategies ...