We thank Anthony J. Koleske, PhD for providing the YFP mice.

Part 1: Microprism Optics and Microscope Setup
<ol>
<li>The microprisms we use are 1-mm, right-angle prisms made from BK7 glass purchased from the Optosigma Corporation (Figure 1). Microprisms are handled using forceps whenever possible.</li>
<li>The hypotenuse of the microprism is coated with enhanced silver to provide a reflectivity of greater than 97% from 400 &#x2013; 2000 nm. This provides internal reflection of both the laser excitation light and the emitted fluorescence.</li>
<li>Microprisms are always handled using forceps when possible. Gloves are worn at all times to keep the prism clean. </li>
<li>We use a custom-built two-photon microscope capable of small-animal imaging. The animal, attached to the stereotactic device, is placed on a 3-axis motorized stage.</li>
<li>The microscope is connected to a Windows PC running ScanImage software. ScanImage is image acquisition software written in Matlab by Pologruto et al1.</li>
<li>For microprism imaging, we collect images at a resolution of 1024 x 1024 or 512 x 512 pixels. In addition, we typically scan at 2 ms/line or 4 ms/line.</li>
<li>Two types of objectives are used in the experiments: 
<ul>
<li>A lower numerical aperture (NA) air objective for wide field-of-view imaging (see Table of Reagents and Equipment).</li>
<li>A higher NA air objective with a glass correction collar capable of minimizing spherical aberrations induced by 0 &#x2013; 2 mm of glass (see Table of Reagents and Equipment).</li>
</ul>
</li>
<li>When the animal is ready for imaging under a low magnification objective, align the square raster scanning pattern of the laser with the top of the microprism. This will help orient the image and make efficient use of the laser power.</li>
<li>Using a higher NA objective with a glass correction collar, set the correction collar to correct for approximately 1 mm of glass. Once a fluorescent image appears during the experiment, one can carefully reach inside the microscope to optimize the correction collar to maximize the imaging resolution.</li>
</ol>
Part 2: Non-survival Animal Surgery and Microprism Insertion
<ol>
<li>The entire procedure length, from the initial incision until completion of imaging, depends on the objectives of the experiment. However, the non-survival surgical procedure and microprism placement can be expected to take approximately 30 &#x2013; 40 minutes to complete.</li>
<li>Mice (P30-P60) are weighed and anesthetized appropriately using an I.P. injection of ketamine (100 mg/kg) and xylazine (10 mg/kg) to a plane suitable for surgical procedures.</li>
<li>Throughout the entire experiment, the animal should be checked every 15-20 minutes to assess its level of anesthesia. If the animal reacts to a firm toe-pinch, a supplemental dose of ketamine/xylazine is required. Typically a supplemental dose is one-third of the initial dose and can be prepared ahead of time in a syringe for immediate I.P. injection if needed.</li>
<li>Once the mouse is properly anesthetized, the majority of the hair on the animal&#x2019;s head is shaved off using a trimmer with a #40 blade.</li>
<li>Nair&reg; is applied to the remaining hairs on the head to help create a surface free of hairs that may get in the way of the microprism during imaging. After 5 minutes, the Nair&reg; is cleaned off using a cotton-tipped swab.</li>
<li>The anesthetized animal is properly placed into a mouse stereotactic device. During the surgical procedures and imaging sessions the animal is kept on a water-filled heating pad set at 36 degrees Celsius.</li>
<li>With the animal&#x2019;s head properly fixed in place, one clean incision is made using a scalpel down the medial line of the skull to separate the skin.</li>
<li>A small amount of 3% hydrogen peroxide is placed on a cotton-tipped swab and applied to the skull to clear away membranes between the skull and the skin.</li>
<li>The hydrogen peroxide also helps to reveal bregma, a major landmark in knowing where to perform the craniotomy and insert the microprism.</li>
<li>Before the craniotomy is started, the ear bars and bite bar are properly adjusted to tilt the exposed skull such that it is basically level with the table. This provides a better platform for the craniotomy. In addition, this makes the top surface of the prism perpendicular with the incoming excitation light. This will help minimize optical aberrations while imaging.</li>
<li>A small dental burr (~0.8 mm head size) is connected to a Dremel&reg; tool using a flexible extension arm. For craniotomies, we set the speed to approximately 7,000 to 10,000 rpms.</li>
<li>The dental burr is skimmed along the skull until a square groove is established in the bone. The sides of the square are approximately 2-3 mm in length and centered 1.5 mm caudal and 1 mm lateral to bregma.</li>
<li>Continuing thinning the bone until a small opening is made through the skull. A pair of forceps can lift the bone flap away from the skull.</li>
<li>The dura needs to be removed before the microprism can be inserted into the cortex. Bleeding is best controlled by letting the blood coagulate on the surface for several minutes. Afterwards, the coagulated blood is removed using a surgical sponge before inserting the microprism.</li>
<li>To help assist the alignment of the prism with the cortex, the vertical face of the microprism is lined up parallel to the handles on the forceps.</li>
<li>With the prism properly held, the entire microprism is inserted until the top of the prism is flush with the neocortical surface.</li>
<li>When inserted correctly, the microprism should remain in the proper position. Any bleeding that results is minimal and can be absorbed with a small surgical sponge. Once the prism is in place the animal is ready for imaging.</li>
<li>After a microprism has been used in an animal, it can be re-used several times. However, it does need to be properly cleaned to remove any remaining biological material. This can be accomplished by dipping the prism into a small volume of HCl acid, followed by a dip into methanol. The cleaned microprism can then be wrapped in lens paper until future use.</li>
</ol>
Part 3: Representative Results
<img src="/files/ftp_upload/1509/1509fig1.jpg" alt="Figure 1" /> Figure 1: Images of the right-angle microprism. Prisms used in this experiment are one-millimeter, BK7 glass prisms with an enhanced silver coating on the hypotenuse to allow internal reflection of excitation and emission light.
<img src="/files/ftp_upload/1509/1509fig2.jpg" alt="Figure 2" /> Figure 2: Image of layer V YFP pyramidal neurons. Wide field-of-view imaging with the microprism reveals a large population of pyramidal cell bodies ~800 &#x2013; 900 &mu;m deep from the cortical surface. In addition, apical dendrites extending up towards layer I can also be resolved. Scale bar = 200 &mu;m.
<img src="/files/ftp_upload/1509/1509fig3.jpg" alt="Figure 3" /> Figure 3: A higher numerical aperture objective used in conjunction with the microprism provides spatial resolution sufficient to resolve dendritic spines on the apical dendrites on layer V pyramidal neurons. Scale bar = 10 &mu;m.
<img src="/files/ftp_upload/1509/1509fig4.jpg" alt="Figure 4" /> Figure 4: An image obtained using the microprism and a tail-vein injection of fluorescent dye. The image shows vertical, large caliber vessels extending from deep brain regions and smaller capillaries branching off through the cortical volume. Scale bar = 200 &mu;m.
<img src="/files/ftp_upload/1509/1509fig5.jpg" alt="Figure 5" /> Figure 5: Using a higher numerical aperture objective, the network of capillaries in the neocortex can be imaged with greater detail. Scale bar = 50 &mu;m.
<img src="/files/ftp_upload/1509/1509fig6.jpg" alt="Figure 6" /> Figure 6: Microprisms also allows for imaging red blood cells (RBCs) flowing through vessels. RBCs do not absorb the fluorescent dye and therefore are seen as dark stripes across the vessel. By line-scanning across the vessel, one can obtain measurements of RBC flux and velocity. Scale bar = 10 &mu;m (horizontal bar in top and bottom image) and 250 ms (vertical bar in bottom image).

<ol>
	<li>Pologruto, T. A., Sabatini, B. L., Svoboda, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ScanImage:+flexible+software+for+operating+laser+scanning+microscopes.">ScanImage: flexible software for operating laser scanning microscopes.</a> Biomed Eng Online. 2, 13-13 (2003).</li><li>Feng, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+neuronal+subsets+in+transgenic+mice+expressing+multiple+spectral+variants+of+GFP.">Imaging neuronal subsets in transgenic mice expressing multiple spectral variants of GFP.</a> Neuron. 28 (1), 41-41 (2000).</li><li>Kleinfeld, D., Mitra, P. P., Helmchen, F., Denk, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluctuations+and+stimulus-induced+changes+in+blood+flow+observed+in+individual+capillaries+in+layers+2+through+4+of+rat+neocortex.">Fluctuations and stimulus-induced changes in blood flow observed in individual capillaries in layers 2 through 4 of rat neocortex.</a> Proc Natl Acad Sci U S A. 95 (26), 15741-15741 (1998).</li></ol>

Experiments on animals were performed in accordance with the guidelines and regulations set forth by Yale University's IACUC.

Using a microprism in an imaging experiment is relatively straightforward and offers many advantages for in vivo studies on the neocortex. Representative examples using this technique include images taken from transgenic mice expressing yellow fluorescent protein in layer V cortical neurons. Using microprisms one can see a collection of large pyramidal cell bodies in layer V, nearly 1 mm below the cortical surface. Also seen are the apical dendrites that extend through all the superficial layers before diverging into ...

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		<th>Comment</th>
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<td>With enhanced silver reflective coating-(R77)</td>
<td>&nbsp;</td>
<td>OptoSigma Corp. (Santa Ana, CA)</td>
<td>055-0010</td>
<td>With enhanced silver reflective coating-(R77)</td>
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<td>Fluorescein-dextran</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>FD70</td>
<td>70 kDa </td>
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<tr>
<td>0.28 NA, 4x air objective</td>
<td>&nbsp;</td>
<td>Olympus</td>
<td>XLFLUOR 4x/340</td>
<td>&nbsp;</td>
<tr>
<td>0.60 NA, 40x air objective</td>
<td>&nbsp;</td>
<td>Olympus</td>
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in vivo imaging of deep cortical layers using a microprism

We present a protocol for in vivo imaging of cortical tissue using a deep-brain imaging probe in the shape of a microprism.  Microprisms are 1-mm in size and have a reflective coating on the hypotenuse to allow internal reflection of excitation and emission light.  The microprism probe simultaneously images multiple cortical layers with a perspective typically seen only in slice preparations.  Images are collected with a large field-of-view (~900 &mu;m).  In addition, we provide details on the non-survival surgical procedure and microscope setup.  Representative results include images of layer V pyramidal neurons from Thy-1 YFP-H mice showing their apical dendrites extending through the superficial cortical layer and extending into tufts.  Resolution was sufficient to image dendritic spines near the soma of layer V neurons.  A tail-vein injection of fluorescent dye reveals the intricate network of blood vessels in the cortex.  Line-scanning of red blood cells (RBCs) flowing through the capillaries reveals RBC velocity and flux rates can be obtained. This novel microprism probe is an elegant, yet powerful new method of visualizing deep cellular structures and cortical function in vivo.

Watch this Scientific Journal Video about In vivo Imaging of Deep Cortical Layers using a Microprism at JoVE.com

In vivo Imaging of Deep Cortical Layers using a Microprism

Right-angle microprisms inserted into the mouse neocortex allows for deep imaging of multiple cortical layers with a viewpoint typically found in slice. One-millimeter microprisms offer a wide field-of-view (~900 &mu;m) and spatial resolutions sufficient to resolve dendritic spines. We demonstrate layer V neuronal imaging and neocortical vascular imaging using microprisms.

In vivo Imaging of Deep Cortical Layers using a Microprism

We present a protocol for in vivo imaging of cortical tissue using a deep-brain imaging probe in the shape of a microprism.  Microprisms are 1-mm in size ...

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11.3K Views.  Yale University. Right-angle microprisms inserted into the mouse neocortex allows for deep imaging of multiple cortical layers with a viewpoint typically found in slice. One-millimeter microprisms offer a wide field-of-view (~900 μm) and spatial resolutions sufficient to resolve dendritic spines. We demonstrate layer V neuronal imaging and neocortical vascular imaging using microprisms.

Video: In vivo Imaging of Deep Cortical Layers using a Microprism

Preparation of Dissociated Mouse Cortical Neuron Cultures

This video will guide you through the process for generating cortical neuronal cultures from late embryo and early postnatal mouse brain. These cultures can be used for a variety of applications including immunocytochemistry, biochemistry, electrophysiology, calcium and sodium imaging, protein and/or RNA isolation. These cultures also provide a platform to study the neuronal development of transgenic animals that carry a late embryonic or postnatal lethal gene mutation. The procedure is relatively straight forward, requires some experience in tissue culture technique and should not take longer than two to three hours if you are properly prepared. Careful separation of the cortical rind from the thalamo-cortical fiber tract will reduce the number of unwanted non-neuronal cells. To increase yields of neuronal cells triturate the pieces of the cortical tissue gently after the enzyme incubation step. This is imperative as it prevents unnecessary injury to cells and premature neuronal cell death. Since these cultures are maintained in the absence of glia feeder cells, they also offer an added advantage of growing cultures enriched in neurons.

This video shows a procedure for generating neuronal cultures from late embryo and early postnatal mouse cortex. These cultures can be used for immunocytochemistry, biochemistry, electrophysiology, calcium and sodium imaging and provide a platform to study the neuronal development of transgenic animals that carry a postnatal lethal gene mutation.

This video will guide you through the process for generating cortical neuronal cultures from late embryo and early postnatal mouse brain. These cultures ...

Calcium Imaging of Cortical Neurons using Fura-2 AM

Calcium imaging is a common technique that is useful for measuring calcium signals in cultured cells. Calcium imaging techniques take advantage of calcium indicator dyes, which are BAPTA-based organic molecules that change their spectral properties in response to the binding of Ca2+ ions. Calcium indicator dyes fall into two categories, ratio-metric dyes like Fura-2 and Indo-1 and single-wavelength dyes like Fluo-4. Ratio-metric dyes change either their excitation or their emission spectra in response to calcium, allowing the concentration of intracellular calcium to be determined from the ratio of fluorescence emission or excitation at distinct wavelengths.  The main advantage of using ratio-metric dyes over single wavelength probes is that the ratio signal is independent of the dye concentration, illumination intensity, and optical path length allowing the concentration of intracellular calcium to be determined independently of these artifacts. One of the most common calcium indicators is Fura-2, which has an emission peak at 505 nM and changes its excitation peak from 340 nm to 380 nm in response to calcium binding.  Here we describe the use of Fura-2 to measure intracellular calcium elevations in neurons and other excitable cells.

Calcium signals play a key role in many cellular processes including gene expression, survival and differentiation. Here we demonstrate how to perform calcium imaging using Fura-2 AM. Calcium imaging is a valuable tool to study the regulation of intracellular calcium in real time and its regulation of signaling cascades.

Calcium imaging is a common technique that is useful for measuring calcium signals in cultured cells. Calcium imaging techniques take advantage of calcium ...

In vivo Bioluminescent Imaging of Mammary Tumors Using IVIS Spectrum

4T1 mouse mammary tumor cells can be implanted sub-cutaneously in nu/nu mice to form palpable tumors in 15 to 20 days. This xenograft tumor model system is valuable for the pre-clinical in vivo evaluation of putative antitumor compounds.
The 4T1 cell line has been engineered to constitutively express the firefly luciferase gene (luc2). When mice carrying 4T1-luc2 tumors are injected with Luciferin the tumors emit a visual light signal that can be monitored using a sensitive optical imaging system like the IVIS Spectrum. The photon flux from the tumor is proportional to the number of light emitting cells and the signal can be measured to monitor tumor growth and development. IVIS is calibrated to enable absolute quantitation of the bioluminescent signal and longitudinal studies can be performed over many months and over several orders of signal magnitude without compromising the quantitative result.
Tumor growth can be monitored for several days by bioluminescence before the tumor size becomes palpable or measurable by traditional physical means. This rapid monitoring can provide insight into early events in tumor development or lead to shorter experimental procedures.
Tumor cell death and necrosis due to hypoxia or drug treatment is indicated early by a reduction in the bioluminescent signal. This cell death might not be accompanied by a reduction in tumor size as measured by physical means. The ability to see early events in tumor necrosis has significant impact on the selection and development of therapeutic agents.
Quantitative imaging of tumor growth using IVIS provides precise quantitation and accelerates the experimental process to generate results.

In vivo Bioluminescent Imaging of Mammary Tumors Using IVIS Spectrum

Mammary tumor cells expressing luciferase are implanted subcutaneously in mice and visualized using optical imaging to monitor tumor growth and development non-invasively in a longitudinal study.

4T1 mouse mammary tumor cells can be implanted sub-cutaneously in nu/nu mice to form palpable tumors in 15 to 20 days. This xenograft tumor model system ...

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Visualizing developing organ formation as well as progession and treatment of disease often heavily relies on the ability to optically interrogate molecular and functional changes in intact living organisms. Most existing optical imaging methods are inadequate for imaging at dimensions that lie between the penetration limits of modern optical microscopy (0.5-1mm) and the diffusion-imposed limits of optical macroscopy (&gt;1cm) [1]. Thus, many important model organisms, e.g. insects, animal embryos or small animal extremities, remain inaccessible for in-vivo optical imaging. 
Although there is increasing interest towards the development of nanometer-resolution optical imaging methods, there have not been many successful efforts in improving the imaging penetration depth. The ability to perform in-vivo imaging beyond microscopy limits is in fact met with the difficulties associated with photon scattering present in tissues. Recent efforts to image entire embryos for example [2,3] require special chemical treatment of the specimen, to clear them from scattering, a procedure that makes them suitable only for post-mortem imaging. These methods however evidence the need for imaging larger specimens than the ones usually allowed by two-photon or confocal microscopy, especially in developmental biology and in drug discovery.
We have developed a new optical imaging technique named Mesoscopic Fluorescence Tomography [4], which appropriate for non-invasive in-vivo imaging at dimensions of 1mm-5mm. The method exchanges resolution for penetration depth, but offers unprecedented tomographic imaging performance and it has been developed to add time as a new dimension in developmental biology observations (and possibly other areas of biological research) by imparting the ability to image the evolution of fluorescence-tagged responses over time. As such it can accelerate studies of morphological or functional dependencies on gene mutations or external stimuli, and can importantly, capture the complete picture of development or tissue function by allowing longitudinal time-lapse visualization of the same, developing organism.
The technique utilizes a modified laboratory microscope and multi-projection illumination to collect data at 360-degree projections. It applies the Fermi simplification to Fokker-Plank solution of the photon transport equation, combined with geometrical optics principles in order to build a realistic inversion scheme suitable for mesoscopic range. This allows in-vivo whole-body visualization of non-transparent three-dimensional structures in samples up to several millimeters in size.
We have demonstrated the in-vivo performance of the technique by imaging three-dimensional structures of developing Drosophila tissues in-vivo and by following the morphogenesis of the wings in the opaque Drosophila pupae in real time over six consecutive hours.

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Mesoscopic fluorescence tomography operates beyond the penetration limits of tissue-sectioning fluorescence microscopy. The technique is based on multi-projection illumination and a photon transport description. We demonstrate in-vivo whole-body 3D visualization of the morphogenesis of GFP-expressing wing imaginal discs in Drosophila melanogaster.

Visualizing  developing organ formation as well as progession and treatment of disease often  heavily relies on the ability to optically interrogate ...

In-vivo Centrifugation of Drosophila Embryos

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in buoyant density. However, when cells are disrupted prior to centrifugation, proteins and organelles in this non-native environment often inappropriately stick to each other. Here we describe a method to separate organelles by density in intact, living Drosophila embryos. Early embryos before cellularization are harvested from population cages, and their outer egg shells are removed by treatment with 50% bleach. Embryos are then transferred to a small agar plate and inserted, posterior end first, into small vertical holes in the agar. The plates containing embedded embryos are centrifuged for 30 min at 3000g. The agar supports the embryos and keeps them in a defined orientation. Afterwards, the embryos are dug out of the agar with a blunt needle. Centrifugation separates major organelles into distinct layers, a stratification easily visible by bright-field microscopy. A number of fluorescent markers are available to confirm successful stratification in living embryos. Proteins associated with certain organelles will be enriched in a particular layer, demonstrating colocalization. Individual layers can be recovered for biochemical analysis or transplantation into donor eggs. This technique is applicable for organelle separation in other large cells, including the eggs and oocytes of diverse species.

In-vivo Centrifugation of Drosophila Embryos

We describe a method to separate organelles by density in living Drosophila embryos. Embryos are embedded in agar and centrifuged. This technique yields reproducible separation of major organelles along the anterior-posterior embryo axis. This method facilitates colocalization experiments and yields organelle fractions for biochemical analysis and transplantation experiments.

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in ...

Imaging Protein-protein Interactions in vivo

Protein-protein interactions are a hallmark of all essential cellular processes. However, many of these interactions are transient, or energetically weak, preventing their identification and analysis through traditional biochemical methods such as co-immunoprecipitation. In this regard, the genetically encodable fluorescent proteins (GFP, RFP, etc.) and their associated overlapping fluorescence spectrum have revolutionized our ability to monitor weak interactions in vivo using F&ouml;rster resonance energy transfer (FRET)1-3. Here, we detail our use of a FRET-based proximity assay for monitoring receptor-receptor interactions on the endothelial cell surface.

Imaging Protein-protein Interactions in vivo

This protocol describes how to image protein-protein interactions using a FRET-based proximity assay.

Protein-protein interactions are a hallmark of all essential cellular processes. However, many of these interactions are transient, or energetically weak, ...

Single-molecule Imaging of Gene Regulation In vivo Using Cotranslational Activation by Cleavage (CoTrAC)

We describe a fluorescence microscopy method, Co-Translational Activation by Cleavage (CoTrAC) to image the production of protein molecules in live cells with single-molecule precision without perturbing the protein's functionality. This method makes it possible to count the numbers of protein molecules produced in one cell during sequential, five-minute time windows. It requires a fluorescence microscope with laser excitation power density of ~0.5 to 1 kW/cm2, which is sufficiently sensitive to detect single fluorescent protein molecules in live cells. The fluorescent reporter used in this method consists of three parts: a membrane targeting sequence, a fast-maturing, yellow fluorescent protein and a protease recognition sequence. The reporter is translationally fused to the N-terminus of a protein of interest. Cells are grown on a temperature-controlled microscope stage. Every five minutes, fluorescent molecules within cells are imaged (and later counted by analyzing fluorescence images) and subsequently photobleached so that only newly translated proteins are counted in the next measurement.
Fluorescence images resulting from this method can be analyzed by detecting fluorescent spots in each image, assigning them to individual cells and then assigning cells to cell lineages. The number of proteins produced within a time window in a given cell is calculated by dividing the integrated fluorescence intensity of spots by the average intensity of single fluorescent molecules. We used this method to measure expression levels in the range of 0-45 molecules in single 5 min time windows. This method enabled us to measure noise in the expression of the &lambda; repressor CI, and has many other potential applications in systems biology.

Single-molecule Imaging of Gene Regulation In vivo Using Cotranslational Activation by Cleavage CoTrAC

We describe a fluorescence microscopy method, Co-Translational Activation by Cleavage (CoTrAC), to image the production of protein molecules in live cells with single-molecule precision without perturbing the protein's functionality. This method has been used to follow the stochastic expression dynamics of a transcription factor, the &lambda; repressor CI 1.

We describe a fluorescence microscopy method, Co-Translational Activation by Cleavage (CoTrAC) to image the production of protein molecules in live cells ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Quantitation of Protein Expression and Co-localization Using Multiplexed Immuno-histochemical Staining and Multispectral Imaging

Immunohistochemistry is a commonly used clinical and research lab detection technique for investigating protein expression and localization within tissues. Many semi-quantitative systems have been developed for scoring expression using immunohistochemistry, but inherent subjectivity limits reproducibility and accuracy of results. Furthermore, the investigation of spatially overlapping biomarkers such as nuclear transcription factors is difficult with current immunohistochemistry techniques. We have developed and optimized a system for simultaneous investigation of multiple proteins using high throughput methods of multiplexed immunohistochemistry and multispectral imaging. Multiplexed immunohistochemistry is performed by sequential application of primary antibodies with secondary antibodies conjugated to horseradish peroxidase or alkaline phosphatase. Different chromogens are used to detect each protein of interest. Stained slides are loaded into an automated slide scanner and a protocol is created for automated image acquisition. A spectral library is created by staining a set of slides with a single chromogen on each. A subset of representative stained images are imported into multispectral imaging software and an algorithm for distinguishing tissue type is created by defining tissue compartments on images. Subcellular compartments are segmented by using hematoxylin counterstain and adjusting the intrinsic algorithm. Thresholding is applied to determine positivity and protein co-localization. The final algorithm is then applied to the entire set of tissues. Resulting data allows the user to evaluate protein expression based on tissue type (ex. epithelia vs. stroma) and subcellular compartment (nucleus vs. cytoplasm vs. plasma membrane). Co-localization analysis allows for investigation of double-positive, double-negative, and single-positive cell types. Combining multispectral imaging with multiplexed immunohistochemistry and automated image acquisition is an objective, high-throughput method for investigation of biomarkers within tissues.

Immunohistochemistry is a powerful lab technique for evaluating protein localization and expression within tissues. Current semi-automated methods for quantitation introduce subjectivity and often create irreproducible results. Herein, we describe methods for multiplexed immunohistochemistry and objective quantitation of protein expression and co-localization using multispectral imaging.

Immunohistochemistry is a commonly used clinical and research lab detection technique for investigating protein expression and localization within ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...