Financial support for this work from the Science Foundation Ireland, grants SFI/12/RC/2276_P2, SFI/17/RC-PhD/3484 and 18/SP/3522, and Breakthrough Cancer Research (Precision Oncology Ireland) is gratefully acknowledged.

All the procedures with animals were performed under authorizations issued by the Health Products Regulatory Authority (HPRA, Ireland) in accordance with the European Communities Council Directive (2010/63/EU) and were approved by the Animal Experimentation Ethics Committee of the University College Cork.
1. Sample preparation
<ol>
	<li>Staining with the probe of live tissue samples ex vivo&#8203;
	<ol>
		<li>For ex vivo applications, use freshly isolated tissue samples from 4 week old female Balb/c mice.</li>
		<li>On the day of the experiment, euthanize a mouse by decapitation, and quickly dissect fragments of the colon (large intestine), approximately 10 mm in size. Wash them immediately with PBS buffer, place in DMEM medium supplemented with 10 mM Hepes buffer (pH 7.2), and incubate at 37 &#176;C36.</li>
		<li>For surface staining of the serosal side of the intestine, transfer the live tissue samples into a mini-dish, apply 2 mL of complete DMEM containing 1 mg/mL NanO2-IR probe to cover the tissue samples, and incubate for 30 min at 37 &#176;C. 
		NOTE: Cells in post-mortem tissue remain live for many hours in culture. NaNO2-IR shows minor cytotoxicity, so all the experiments were completed within 4 h after tissue isolation.</li>
		<li>For deep tissue intraluminal ex vivo staining, transfer the pieces of the intestine to a dry Petri dish, and remove any excess DMEM with filter paper.</li>
		<li>Inject 1 &#181;L of DMEM containing 1 mg/mL NanO2-IR35 into the lumen with a Hamilton syringe, and incubate the samples for 15 min or for up to 4 h. 
		NOTE: NaNO2-IR shows minor long-term cytotoxic effects; therefore, all the experiments should be completed within 4 h after tissue isolation.</li>
	</ol>
	</li>
	<li>Preparation of stained tumor tissue in live animals
	<ol>
		<li>For in vivo applications, pre-stain CT26 cells for 18 h in serum-free medium containing NaNO2-IR probe at 0.05 mg/mL.</li>
		<li>Take a mouse, shave the area of injection in the right flank, and inject with a syringe 200 &#181;L of a mixture of 1 &#215; 105 non-stained cells and 1 &#215; 105 cells pre-stained with NanO2-IR .</li>
		<li>Allow tumors to grow in the mice, monitoring the tumor size with a caliper and the animal weight periodically37. The animals with grafted tumors become ready for imaging on the seventh day of tumor growth. 
		NOTE: The tumor volume was calculated using equation (1): 
		V = (L &#215; W2)/2&#160; &#160; &#160;(1) 
		&#8203;where L is the diameter of the tumor, and W is the diameter perpendicular to the diameter L.</li>
		<li>Sacrifice the animals by cervical dislocation just before the imaging.</li>
	</ol>
	</li>
</ol>
2. PLIM imaging setup
<ol>
	<li>Take the Cricket adaptor, and remove its back side C-mount adapter to gain access to the intensifier housing inside. Insert the MCP-125 image intensifier into this compartment, and put the C-mount adaptor back.</li>
	<li>Remove the Cricket&#39;s front side C-mount adaptor, insert the 760 nm &#177; 50 nm emission filter, and fix it by putting back the C-mount.</li>
	<li>Connect the Tpx3Cam camera module to the back side of the Cricket module via its C-mount adaptor</li>
	<li>Connect the lens to the front side of the Cricket module via its C-mount adaptor.</li>
	<li>Mount the whole camera assembly on top of the optical black box, facing down to the stage on which the samples will be imaged (Figure 1).</li>
	<li>Mount the 624 nm super-bright LED on a post connected to a breadboard inside the black box.</li>
	<li>Connect the LED to a power supply and a pulse generator. Switch on the LED, and focus it to ensure effective and uniform excitation of imaged samples.</li>
	<li>Connect the camera to another pulse generator, and synchronize the pulses sent to the camera and the LED38.</li>
	<li>Using the special cable and socket on the Cricket unit, connect the intensifier to a standard power supply, and set the gain to 2.7 V.</li>
	<li>Using the focusing capabilities of the lens and Cricket adaptor, focus the camera optics on the sample stage to generate clear images of samples with good contrast and brightness.</li>
	<li>For imager use with Pt-porphyrin dyes, replace the 625 nm LED with a 390 nm LED for excitation, and replace the 760 nm &#177; 50 nm filter with a 650 nm &#177; 50 nm filter in the Cricket module.</li>
</ol>
3. Image acquisition
<ol>
	<li>Place the sample in front of the camera lens. 
	NOTE: Use an x-y-z adjustable stage as the sample holder in order to adjust the sample position for good focus.</li>
	<li>Turn off all the lights in the room.</li>
	<li>Switch on the Sophy software for tuning the operational parameters, such as the focusing and sample alignment. 
	NOTE: Sophy software is provided together with the camera to set up the imaging parameters and record the data. Make sure that the software is connected to the camera by checking the camera code. However, we used a different program for data acquisition.</li>
	<li>In Modules, select infinite frames, and set the pixel operation mode to time over threshold.</li>
	<li>In Modules, go to Preview, and select Active module. This opens the Medpix/Timpix Frames window.</li>
	<li>In this window, change the color scale, and rotate the image to the desired orientation.</li>
	<li>Switch on the intensifier, and start the recording. 
	NOTE: Use the recording screen of the Sophy software to visually confirm the alignment and focus of the sample and to optimize the LED excitation parameters for recording.</li>
	<li>Stop the recording, and close the Sophy software.</li>
	<li>Go to the terminal, and use the custom-designed software to acquire the raw data in the binary format and post-process it (https://github.com/svihra/TimePix3).
	<ol>
		<li>In the terminal, run the following commands to record the data: 
		Cd Document/SPIDR/trunk/Release/ 
		&#160;ls 
		./Tpx3daq - i 1 - b 50 - m - s {Name of the file} - t {acquisition time} 
		NOTE: Upon typing &#34;ls,&#34; check the list of files in the current directory; confirm that Tpx3daq is seen.</li>
		<li>Wait until all the frames are recorded.</li>
		<li>To process the data, run the following commands in the terminal: 
		cd Documents/DataProcessing/Timepix3/Timepix3/ 
		root 
		.L dataprocess.cpp++ 
		&#8203;.x DRGUI.C</li>
		<li>Wait for an RRGui window to open. Select all the variables on the left and All Data, Single File, and Centroid on the right.</li>
		<li>Select the file to process and run the data reducer. 
		&#8203;NOTE: All the processed files will appear in the same folder as the raw file.</li>
	</ol>
	</li>
</ol>
4. Data analysis
<ol>
	<li>Analyze the post-processed data with a dedicated program written in C-language that will write the data into an .ics image file (https://github.com/lmhirvonen/timepix3cam).</li>
	<li>Open the .ics image files using the freely available Time Resolved Imaging software (see Table of Materials). Use two-exponential functions to fit the phosphorescence decays.</li>
	<li>Open the fitted .ics image files with the available image analysis software (see Table of Materials).</li>
	<li>Using Lookup Tables, generate phosphorescence lifetime images, and encode them in pseudocolor scale (e.g., blue color for short lifetimes and red for long lifetimes). Use the Measure function to calculate the average lifetime values for the entire image or specific regions of interest (ROIs).</li>
	<li>Convert the lifetime values into the oxygen concentration by using the equation obtained from the fitting of the O2 calibration of the probe36. 
	NOTE: Equation (2) was used for this work: 
	O2 [&#181;M] = &#8722;86.16 + 770.35 &#215; e&#8722;0.049 &#215; LT&#160; &#160; &#160;(2)</li>
</ol>

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The authors have no conflicts of interest to declare.

The above protocols give a detailed description of the assembly of the new imager and its operation in the microsecond FLIM/PLIM mode. The TCSPC-based new generation Tpx3Cam camera, coupled by means of the opto-mechanical adaptor Cricket with the image intensifier, emission filter, and macro-lens, produces a stable, compact, and flexible optical module that is easy to operate. The imager was shown to perform well with a range of different samples and analytical tasks, which included the characterization of phosphorescent...

O2 is one of the key environmental parameters for living systems, and knowledge of the distribution of O2 and its dynamics is important for many biological studies1,2,3. The assessment of tissue oxygenation by means of phosphorescent probes4,5,6,7,8 and PLIM9,10,11,12,13 are gaining popularity in biological and medical research3,9,14,15,16,17,18,19. This is because PLIM, unlike fluorescence or phosphorescence intensity measurements, is not affected by external factors such as probe concentration, photobleaching, excitation intensity, optical alignment, scattering, and autofluorescence.
However, current O2 PLIM platforms are limited by their sensitivity, image acquisition speed, accuracy, and general usability. Time-correlated single photon counting (TCSPC), combined with a raster scanning procedure, is frequently used in PLIM and fluorescence lifetime imaging microscopy (FLIM) devices20,21,22. However, since PLIM requires a long pixel dwell time (in the millisecond range), the time of image acquisition is much longer than what is required for FLIM applications20,22,23. Other techniques, such as gated CCD/CMOS cameras, lack single photon sensitivity and have low frame rates20,24,25,26. Moreover, the existing PLIM systems are mostly used in the microscopic format, while macroscopic systems are less common27.
The TCSPC-based PLIM macro imager28 was set up to overcome many of these limitations. The design of the imager was greatly facilitated by the use of a new opto-mechanical adapter, Cricket, which has the following: i) two C-mount adapters, which provide easy coupling of the camera module on the back side and objective lens on the front side; ii) an internal housing for an image intensifier and a power socket for the latter on the outer side of the Cricket; iii) an internal space behind the front-side C-mount adapter where a standard 25 mm emission filter can be housed in front of the intensifier; and iv) a built-in light collimating optics with ring regulators, which allow optical alignment/focusing between the lens and the camera to produce crisp images on the camera chip.
In the assembled imager, the camera module is coupled to the back side of the Cricket adapter, which also houses an image intensifier consisting of a photocathode followed by a microchannel plate (MCP), an amplifier, and a fast scintillator, P47 phosphor. A 760 nm &#177; 50 nm emission filter is fitted inside the Cricket, and an objective lens, NMV-50M11&#39;&#39;, is attached to the front side C-mount adapter. Finally, the lens and the camera are aligned optically with ring regulators.
The role of the intensifier is to detect incoming photons and convert them into fast bursts of light on the camera chip, which are registered and used to generate emission decays and lifetime images. The camera module comprises an advanced TCSPC-based optical sensor array (256 pixels x 256 pixels) and a new generation readout chip29,30,31,32,33, which allow the simultaneous recording of the time of arrival (TOA) and the time over threshold (TOT) of photon bursts at each pixel of the imaging chip with a time resolution of 1.6 ns and an 80 Mpixel/s readout rate.
In this configuration, the camera with the intensifier has single-photon sensitivity. It is data-driven and based on the speedy pixel detector readout (SPIDR) system34. The spatial resolution of the imager was previously characterized with planar phosphorescent O2 sensors and a resolution plate mask. The instrument response function (IRF) was measured by the imaging of a planar fluorescent sensor under the same settings as used for all the other measurements. The lifetime of the dye of around 2.6 ns was short enough for it to be used for the IRF measurement in PLIM mode. The imager can image objects of up to 18 mm x 18 mm in size with spatial and temporal resolutions of 39.4 &#181;m and 30.6 ns (full width at half-maximum), respectively28.
The following protocols describe the assembly of the macro imager and its subsequent use for mapping the O2 concentration in biological samples stained with the previously characterized near-infrared O2 probe, NanO2-IR35. The probe is a bright, photostable, cell-permeable O2-sensing probe based on platinum (II) benzoporphyrin (PtBP) dye. It is excitable at 625 nm, emits at 760 nm, and provides a robust optical response to O2 in the physiological range (0%-21% or 0-210 &#181;M of O2). The imager is also demonstrated to characterize different sensor materials based on Pt(II)-porphyrin dyes. Overall, the imager is compact and flexible, similar to a common photographic camera. In the current setup, the imager is appropriate for different widefield PLIM applications. Substituting the LED with a fast laser source will further improve the performance of the imager and could potentially enable nanosecond FLIM applications.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>627 nm LED</td>
			<td>Parts Express</td>
			<td></td>
			<td>Can be replaced with different LED based on the excitation wavelength of the sensor. Used 390 nm LED for Pt-porphyrin dyes.</td>
		</tr>
		<tr>
			<td>760 &#177; 50 nm emission filter</td>
			<td>Edmund Optics</td>
			<td>84-788</td>
			<td>Can be replaced with different filter based on the emission wavelength of the sensor. Used 650 &#177; 50 nm bandpass filter for Pt-porphyrin dyes.</td>
		</tr>
		<tr>
			<td>Balb/c mice</td>
			<td>Envigo, UK</td>
			<td>Balb/c</td>
			<td></td>
		</tr>
		<tr>
			<td>Black box</td>
			<td>Thorlabs</td>
			<td>XE25C9/M</td>
			<td></td>
		</tr>
		<tr>
			<td>Cricket Adapter</td>
			<td>Photonis</td>
			<td>Cricket-2</td>
			<td></td>
		</tr>
		<tr>
			<td>CT26 cells&#160;</td>
			<td>ATCC</td>
			<td>CT26.WT</td>
			<td>https://www.atcc.org/products/crl-2638</td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Sigma-Aldrich</td>
			<td>D0697</td>
			<td>Other media can also be used</td>
		</tr>
		<tr>
			<td>ImageJ Software</td>
			<td>ImageJ</td>
			<td></td>
			<td>Free Image analysis software. Can be downloaded from: https://imagej.nih.gov/ij/index.html</td>
		</tr>
		<tr>
			<td>MCP-125 image intensifier with P47 phosphor screen</td>
			<td>Photonis</td>
			<td>PP0360EF</td>
			<td></td>
		</tr>
		<tr>
			<td>Mini dishes</td>
			<td>Sarstedt</td>
			<td>83.3900.300</td>
			<td>35 mm diameter&#160;</td>
		</tr>
		<tr>
			<td>Mylar plastic film, 75 micron&#160;</td>
			<td>RS Ireland</td>
			<td>785-0795</td>
			<td>Othe plastic substrates can also be used</td>
		</tr>
		<tr>
			<td>NanO2-IR</td>
			<td>home-made</td>
			<td>n/a</td>
			<td>The probe can be synthesised according to the published method &#39;Tsytsarev V, Arakawa H, Borisov S, Pumbo E, Erzurumlu RS, Papkovsky DB. In vivo imaging of brain metabolism activity using a phosphorescent oxygen-sensitive probe. J Neurosci Methods. 2013 Jun 15;216(2):146-51. doi: 10.1016/j.jneumeth.2013.04.005. Epub 2013 Apr 25. PMID: 23624034; PMCID: PMC3719178.&#39; or provided by our lab.&#160;</td>
		</tr>
		<tr>
			<td>NMV-50M11&#8221; 50 mm lens</td>
			<td>Navitar</td>
			<td></td>
			<td>Other lenses compatibel with C-mount adators can be used</td>
		</tr>
		<tr>
			<td>Optical breadboard</td>
			<td>Thorlabs</td>
			<td>MB1836</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri Dishes</td>
			<td>Sarstedt</td>
			<td>82.1472.001</td>
			<td>92 mm diameter</td>
		</tr>
		<tr>
			<td>Power Supply</td>
			<td>Tenma</td>
			<td>72-10495</td>
			<td></td>
		</tr>
		<tr>
			<td>Pulse Generator</td>
			<td>Tenma</td>
			<td>TGP110</td>
			<td></td>
		</tr>
		<tr>
			<td>Sophy</td>
			<td>Amsterdam Scientific Instruments</td>
			<td>n/z</td>
			<td>Provided by ASI together with the Tpx3Cam</td>
		</tr>
		<tr>
			<td>Tpx3Cam</td>
			<td>Amsterdam Scientific Instruments</td>
			<td>TPXCAM</td>
			<td></td>
		</tr>
		<tr>
			<td>Tri2 Software</td>
			<td>University of Oxford</td>
			<td>n/a</td>
			<td>Free Time Resolved Imaging software, can be downloaded from: https://users.ox.ac.uk/~atdgroup/index.shtml</td>
		</tr>
		<tr>
			<td>XYZ Translation Stage</td>
			<td>Thorlabs</td>
			<td>LT3</td>
			<td></td>
		</tr>
	</tbody>
</table>

fluorescence lifetime macro imager for biomedical applications

This paper presents a new photoluminescence lifetime imager designed to map the molecular oxygen (O2) concentration in different phosphorescent samples ranging from solid-state, O2-sensitive coatings to live animal tissue samples stained with soluble O2-sensitive probes. In particular, the nanoparticle-based near-infrared probe NanO2-IR, which is excitable with a 625 nm light-emitting diode (LED) and emits at 760 nm, was used. The imaging system is based on the Timepix3 camera (Tpx3Cam) and the opto-mechanical adaptor, which also houses an image intensifier. O2 phosphorescence lifetime imaging microscopy (PLIM) is commonly required for various studies, but current platforms have limitations in their accuracy, general flexibility, and usability.
The system presented here is a fast and highly sensitive imager, which is built on an integrated optical sensor and readout chip module, Tpx3Cam. It is shown to produce high-intensity phosphorescence signals and stable lifetime values from surface-stained intestinal tissue samples or intraluminally stained fragments of the large intestine and allows the detailed mapping of tissue O2 levels in about 20 s or less. Initial experiments on the imaging of hypoxia in grafted tumors in unconscious animals are also presented. We also describe how the imager can be re-configured for use with O2-sensitive materials based on Pt-porphyrin dyes using a 390 nm LED for the excitation and a bandpass 650 nm filter for emission. Overall, the PLIM imager was found to produce accurate quantitative measurements of lifetime values for the probes used and respective two-dimensional maps of the O2 concentration. It is also useful for the metabolic imaging of ex vivo tissue models and live animals.

For ex vivo imaging applications, fragments of intestinal tissues were stained by the topical application of the NanO2-IR probe on the serosal side of the tissue. For deeper staining, 1 &#181;L of the probe was injected into the lumen. In the latter case, the 0.2-0.25 mm thick intestinal wall shielded the probe from the camera. The two staining processes are demonstrated in Figure 2A.
The resulting intensity and PLIM images are presented in <strong class=...

Watch this Scientific Journal Video about Fluorescence Lifetime Macro Imager for Biomedical Applications at JoVE.com

Fluorescence Lifetime Macro Imager for Biomedical Applications

This paper describes the use of a new, fast optical imager for the macroscopic photoluminescence lifetime imaging of long decay emitting samples. The integration, image acquisition, and analysis procedures are described, along with the preparation and characterization of the sensor materials for the imaging and the application of the imager in studying biological samples.

This paper presents a new photoluminescence lifetime imager designed to map the molecular oxygen (O2) concentration in different phosphorescent samples ...

fluorescence-lifetime-macro-imager-for-biomedical-applications

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JoVE Journal

Biology

661 Views.  University College Cork. This paper describes the use of a new, fast optical imager for the macroscopic photoluminescence lifetime imaging of long decay emitting samples. The integration, image acquisition, and analysis procedures are described, along with the preparation and characterization of the sensor materials for the imaging and the application of the imager in studying biological samples.

Video: Fluorescence Lifetime Macro Imager for Biomedical Applications

BioMEMS and Cellular Biology: Perspectives and Applications

The ability to culture cells has revolutionized hypothesis testing in basic cell and molecular biology research. It has become a standard methodology in drug screening, toxicology, and clinical assays, and is increasingly used in regenerative medicine. However, the traditional cell culture methodology   essentially consisting of the immersion of a large population of cells in a homogeneous fluid medium and on a homogeneous flat substrate   has become increasingly limiting both from a fundamental and practical perspective. Microfabrication technologies have enabled researchers to design, with micrometer control, the biochemical composition and topology of the substrate, and the medium composition, as well as the neighboring cell type in the surrounding cellular microenvironment. Additionally, microtechnology is conceptually well-suited for the development of fast, low-cost in vitro systems that allow for high-throughput culturing and analysis of cells under large numbers of conditions. In this interview, Albert Folch explains these limitations, how they can be overcome with soft lithography and microfluidics, and describes some relevant examples of research in his lab and future directions.

The ability to culture cells has revolutionized hypothesis testing in basic cell and molecular biology research. It has become a standard methodology in ...

Microfluidic Applications for Disposable Diagnostics

In this interview, Dr. Klapperich discusses the fabrication of thermoplastic microfluidic devices and their application for development of new diagnostics.

In this interview, Dr. Klapperich discusses the fabrication of thermoplastic microfluidic devices and their application for development of new diagnostics.

In this interview, Dr. Klapperich discusses the fabrication of thermoplastic microfluidic devices and their application for development of new ...

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

Fluorescence Resonance Energy Transfer (FRET) microscopy has been widely used to study the structure and dynamics of molecules of biological interest, such as nucleic acids and proteins. Single molecule FRET (sm-FRET) measurements on immobilized molecules permit long observations of the system -effectively until both dyes photobleach- resulting in time-traces that report on biomolecular dynamics with a broad range of timescales from milliseconds to minutes. To facilitate the acquisition of large number of traces for statistical analyses, the process must be automated and the sample environment should be tightly controlled over the entire measurement time (~12 hours). This is accomplished using an automated scanning confocal microscope that allows the interrogation of thousands of single molecules overnight, and a microfluidic cell that permits the controlled exchange of buffer, with restricted oxygen content and maintains a constant temperature throughout the entire measuring period. Here we show how to assemble the microfluidic device and how to activate its surface for DNA immobilization. Then we explain how to prepare a buffer to maximize the photostability and lifetime of the fluorophores. Finally, we show the steps involved in preparing the setup for the automated acquisition of time-resolved single molecule FRET traces of DNA molecules.

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

Fluorescence Resonance Energy Transfer (FRET) microscopy has been widely used to study the structure and dynamics of molecules of biological interest, ...

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

An ex-ovo Chicken Embryo Culture System Suitable for Imaging and Microsurgery Applications

Understanding the relationships between genetic and microenvironmental factors that drive normal and malformed embryonic development is fundamental for discovering new therapeutic strategies. Advancements in imaging technology have enabled quantitative investigation of the organization and maturing of the body plan, but later stage embryonic morphogenesis is less clear. Chicken embryos are an attractive vertebrate animal model system for this application because of its ease of culture and surgical manipulation. Early embryos can be cultured for a short time on filter paper rings, which enables complete optical access for cell patterning and fate studies1,2. Studying advanced developmental processes such as cardiac morphogenesis are traditionally performed through a window of the eggshell3-5, but this technique limits optical access due to window size. We previously developed a simple method to culture whole embryos ex-ovo on hexagonal weigh boats for up to 10 days, which enabled high resolution imaging via ultrasonography6,7. These cultures were difficult to transport, limiting the types of imaging tools available for live experiments. We here present an improved shell-less culture system with a cost-effective, portable environmental chamber. Eggs were cracked onto a hammock created by a polyurethane membrane (cling wrap) affixed circumferentially to a plastic cup partially filled with sterile water. The dimensions of the circumference and depth of the hammock were both critical to maintain surface tension, while the mechanics of the hammock and water beneath helped dampen vibrations induced by transportation. A small footprint circulating water bath was also developed to enable continuous temperature control during experimentation. We demonstrate the ability to culture embryos in this way for at least 14 days without morphogenic defect or delay and employ this system in several microsurgical and imaging applications.

An ex-ovo Chicken Embryo Culture System Suitable for Imaging and Microsurgery Applications

In this article, we present a simple methodology to enable long-term ex-ovo avian embryo culture. This technique is ideal for longitudinal experimentation requiring complete optical accessibility and/or sterile transportation in avian embryos.

Understanding the relationships between genetic and microenvironmental factors that drive normal and malformed embryonic development is fundamental for ...

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a quality control mechanism that can target and selectively degrade cellular material including organelles1-4. For example, damaged or redundant mitochondria (Fig. 1), not disposed of by autophagy, can represent a threat to cellular homeostasis and cell survival. In the yeast, Saccharomyces cerevisiae, nutrient deprivation (e.g., nitrogen starvation) or damage can promote selective turnover of mitochondria by autophagy in a process termed mitophagy 5-9.
We describe a simple fluorescence microscopy approach to assess autophagy. For clarity we restrict our description here to show how the approach can be used to monitor mitophagy in yeast cells. The assay makes use of a fluorescent reporter, Rosella, which is a dual-emission biosensor comprising a relatively pH-stable red fluorescent protein linked to a pH-sensitive green fluorescent protein. The operation of this reporter relies on differences in pH between the vacuole (pH ~ 5.0-5.5) and mitochondria (pH ~ 8.2) in living cells. Under growing conditions, wild type cells exhibit both red and green fluorescence distributed in a manner characteristic of the mitochondria. Fluorescence emission is not associated with the vacuole. When subjected to nitrogen starvation, a condition which induces mitophagy, in addition to red and green fluorescence labeling the mitochondria, cells exhibit the accumulation of red, but not green fluorescence, in the acidic vacuolar lumen representing the delivery of mitochondria to the vacuole. Scoring cells with red, but not green fluorescent vacuoles can be used as a measure of mitophagic activity in cells5,10-12.

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

A robust approach to monitor the delivery of organelles to the acidic lumen of the yeast vacuole for degradation and recycling is described. The method relies on the specific labeling of target organelles with a genetically encoded dual-emission fluorescence pH-biosensor, and visualization of individual cells using fluorescence microscopy.

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a ...

Absolute Quantum Yield Measurement of Powder Samples

Measurement of fluorescence quantum yield has become an important tool in the search for new solutions in the development, evaluation, quality control and research of illumination, AV equipment, organic EL material, films, filters and fluorescent probes for bio-industry.
Quantum yield is calculated as the ratio of the number of photons absorbed, to the number of photons emitted by a material. The higher the quantum yield, the better the efficiency of the fluorescent material.
For the measurements featured in this video, we will use the Hitachi F-7000 fluorescence spectrophotometer equipped with the Quantum Yield measuring accessory and Report Generator program. All the information provided applies to this system. 
Measurement of quantum yield in powder samples is performed following these steps:
<ol>
 <li>Generation of instrument correction factors for the excitation and emission monochromators. This is an important requirement for the correct measurement of quantum yield. It has been performed in advance for the full measurement range of the instrument and will not be shown in this video due to time limitations.</li>
 <li>Measurement of integrating sphere correction factors. The purpose of this step is to take into consideration reflectivity characteristics of the integrating sphere used for the measurements.</li>
 <li>Reference and Sample measurement using direct excitation and indirect excitation.</li>
 <li>Quantum Yield calculation using Direct and Indirect excitation. Direct excitation is when the sample is facing directly the excitation beam, which would be the normal measurement setup. However, because we use an integrating sphere, a portion of the emitted photons resulting from the sample fluorescence are reflected by the integrating sphere and will re-excite the sample, so we need to take into consideration indirect excitation. This is accomplished by measuring the sample placed in the port facing the emission monochromator, calculating indirect quantum yield and correcting the direct quantum yield calculation.</li>
 <li>Corrected quantum yield calculation.</li>
 <li>Chromaticity coordinates calculation using Report Generator program.</li>
</ol>
The Hitachi F-7000 Quantum Yield Measurement System offer advantages for this
application, as follows:
<ul>
 <li>High sensitivity (S/N ratio 800 or better RMS). Signal is the Raman band of water measured under the following conditions: Ex wavelength 350 nm, band pass Ex and Em 5 nm, response 2 sec), noise is measured at the maximum of the Raman peak. High sensitivity allows measurement of samples even with low quantum yield. Using this system we have measured quantum yields as low as 0.1 for a sample of salicylic acid and as high as 0.8 for a sample of magnesium tungstate.</li>
 <li>Highly accurate measurement with a dynamic range of 6 orders of magnitude allows for measurements of both sharp scattering peaks with high intensity, as well as broad fluorescence peaks of low intensity under the same conditions. </li>
 <li>High measuring throughput and reduced light exposure to the sample, due to a high scanning speed of up to 60,000 nm/minute and automatic shutter function. </li>
 <li>Measurement of quantum yield over a wide wavelength range from 240 to 800 nm.</li>
 <li>Accurate quantum yield measurements are the result of collecting instrument spectral response and integrating sphere correction factors before measuring the sample.</li>
 <li>Large selection of calculated parameters provided by dedicated and easy to use software.
</li>
</ul> 
During this video we will measure sodium salicylate in powder form which is known to have a quantum yield value of 0.4 to 0.5.

In this video we will demonstrate measuring and calculating absolute quantum yield and chromaticity coordinates directly in powder samples using the Hitachi F-7000 Quantum Yield Measuring System.

Measurement of fluorescence quantum yield has become an important tool in the search for new solutions in the development, evaluation, quality control and ...

Infinium Assay for Large-scale SNP Genotyping Applications

Genotyping variants in the human genome has proven to be an efficient method to identify genetic associations with phenotypes. The distribution of variants within families or populations can facilitate identification of the genetic factors of disease. Illumina&#39;s panel of genotyping BeadChips allows investigators to genotype thousands or millions of single nucleotide polymorphisms (SNPs) or to analyze other genomic variants, such as copy number, across a large number of DNA samples. These SNPs can be spread throughout the genome or targeted in specific regions in order to maximize potential discovery. The Infinium assay has been optimized to yield high-quality, accurate results quickly. With proper setup, a single technician can process from a few hundred to over a thousand DNA samples per week, depending on the type of array. This assay guides users through every step, starting with genomic DNA and ending with the scanning of the array. Using propriety reagents, samples are amplified, fragmented, precipitated, resuspended, hybridized to the chip, extended by a single base, stained, and scanned on either an iScan or Hi Scan high-resolution optical imaging system. One overnight step is required to amplify the DNA. The DNA is denatured and isothermally amplified by whole-genome amplification; therefore, no PCR is required. Samples are hybridized to the arrays during a second overnight step. By the third day, the samples are ready to be scanned and analyzed. Amplified DNA may be stockpiled in large quantities, allowing bead arrays to be processed every day of the week, thereby maximizing throughput.

A protocol is described that uses Illumina&#39;s Infinium assays to perform large-scale genotyping. These assays can reliably genotype millions of SNPs across hundreds of individual DNA samples in three days. Once generated, these genotypes can be used to check for associations with a variety of different diseases or phenotypes.

Genotyping variants in the human genome has proven to be an efficient method to identify genetic associations with phenotypes. The distribution of ...

Recovery of Adult Zebrafish Hearts for High-throughput Applications

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and purification of RNA, DNA, and proteins. All of these applications demand the rapid recovery of significant numbers of zebrafish hearts to avoid gene regulatory, metabolic, and other changes that begin after death. Adult zebrafish hearts are also required for studying heart structure for a variety of mutants and for studying heart regeneration. However, the traditional zebrafish heart dissection is slow and difficult and requires specialized tools, making large-scale dissection of adult zebrafish hearts tedious. Traditional methods also harbor the risk of damaging the heart during the dissection. Here, we describe a method for dissection of adult zebrafish hearts that is fast, reproducible, and preserves heart architecture. Furthermore, this method does not require specialized tools, is painless for the zebrafish, can be performed on fresh or fixed specimens, and can be performed on zebrafish as young as one month old. The approach described expands the use of adult zebrafish for cardiovascular research.

Use of zebrafish for cardiovascular research is expanding towards research on adult hearts. For these applications, quick and simple isolation of cardiac tissues is key to avoid post-mortem changes and to obtain an adequate number of samples. Here, we describe a fast and reproducible method for dissecting adult zebrafish hearts.

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and ...

Genetic Barcoding with Fluorescent Proteins for Multiplexed Applications

Fluorescent proteins, fluorescent dyes and fluorophores in general have revolutionized the field of molecular cell biology. In particular, the discovery of fluorescent proteins and their genes have enabled the engineering of protein fusions for localization, the analysis of transcriptional activation and translation of proteins of interest, or the general tracking of individual cells and cell populations. The use of fluorescent protein genes in combination with retroviral technology has further allowed the expression of these proteins in mammalian cells in a stable and reliable manner. Shown here is how one can utilize these genes to give cells within a population of cells their own biosignature. As the biosignature is achieved with retroviral technology, cells are barcoded &#180;indefinitely&#180;. As such, they can be individually tracked within a mixture of barcoded cells and utilized in more complex biological applications. The tracking of distinct populations in a mixture of cells is ideal for multiplexed applications such as discovery of drugs against a multitude of targets or the activation profile of different promoters. The protocol describes how to elegantly develop and amplify barcoded mammalian cells with distinct genetic fluorescent markers, and how to use several markers at once or one marker at different intensities. Finally, the protocol describes how the cells can be further utilized in combination with cell-based assays to increase the power of analysis through multiplexing.

Since the discovery of the green fluorescent protein gene, fluorescent proteins have impacted molecular cell biology. This protocol describes how expression of distinct fluorescent proteins through genetic engineering is used for barcoding individual cells. The procedure enables tracking distinct populations in a cell mixture, which is ideal for multiplexed applications.

Fluorescent proteins, fluorescent dyes and fluorophores in general have revolutionized the field of molecular cell biology. In particular, the discovery ...