&nbsp;Solutions
<ul>
<li>Intracellular solution for ganglion cells (in mM): 100 K-gluconate, 8 KCl, 1 MgCl2, 1 EGTA, 10 HEPES, 4 ATP, .5 GTP, 90 &micro;M Sulforhodamine B (for staining), adjusted to pH = 7.4 with KOH </li>
<li>Intracellular solution for bipolar cells (in mM): 90 K-gluconate, 8 KCl , 1 MgCl2, 10 BAPTA, 10 HEPES, 4 ATP, .5 GTP, 90 &micro;M Sulforhodamine&nbsp; B (for staining), adjusted to pH = 7.4 with KOH </li>
<li>Extracellular Ringer&rsquo;s solution (in mM): 112 NaCl, 5 Glucose, 5 HEPES, 2 KCl, 2 CaCl2, 1 MgCl2, adjusted to pH = 7.75 with NaOH. Oxygenation of the solution is not necessary. </li>
</ul>
Prepare recording chambers
Our recording chamber consists of a microscope slide with an adhesive well in which small silicone blocks cut out of another adhesive well are placed to provide stability for the tissue slices.
Preparation of retinal slices
To avoid bleaching of the photoreceptors, the dissection is carried out in IR or dim red light light. If one is interested in recording responses from the rod pathway, only IR light should be used (IR dissecting microscope and no room light. Additional IR goggles should be used for procedures not done under the microscope). If one is mainly interested in recording cone light responses, dim red room light or a red flash light can be used. The microscope, however, should still have IR oculars.
If photoreceptor input is irrelevant for the experiment, procedures can be carried out in normal room light using a regular microscope.
<ol>
<li>Prepare two microscope slides with an adhesive silicone well: create two parallel bands of silicone grease (about .5 cm apart) on each slide and place a rectangular piece of filter paper on one side of the slide, crossing both grease bands (push the filter paper with part of a razor blade so that it is well attached to the slide)</li>
<li>Sacrifice a salamander according to established protocols: the animal is placed in an ice bath for 30 minutes, then decapitated and double-pithed.</li>
<li>Remove eye from salamander head and place it under the dissecting microscope on a piece of tissue paper moistened with ice-cold Ringer&rsquo;s solution</li>
<li>Remove connective tissue from the eye</li>
<li>Use razor blade to make a small cut in the cornea</li>
<li>Hold cut cornea with forceps while removing it with scissors along its attachment to the eye</li>
<li>Remove the lens with forceps</li>
<li>Hold the iris with forceps and cut along the ora serrata to remove the iris</li>
<li>Cut the eyecup into two rectangular pieces</li>
<li>Pick up one of the pieces and place it sclera side up on the filter paper (make sure to flatten out the piece while pushing it down very carefully)</li>
<li>Gently lift the sclera off the retina, which now should be attached to the filer paper</li>
<li>Quickly add ice-cold Ringer&rsquo;s solution to the well</li>
<li>Repeat the same for the second piece</li>
<li>Use a razor blade placed in a custom made slicer to cut 200-300&micro;m wide slices. </li>
<li>Use a pair of forceps to pick up the slices at the side of the filter paper, turn them over so the layers of the retina become visible, and place them on the vacuum grease bands</li>
<li>Pick a slice that looks undamaged and shows horizontal bands indicating the layers of the retina (the layers are sometimes difficult to see under the dissection microscope as the magnification is not high enough, however if one can recognize some layering the slice is usually good enough. The final decision on whether to use the slice, however, is made once it is viewed under the upright microscope). Move it to the recording chamber slide by building a Ringer bridge between the two slides</li>
<li>Make sure the filter paper is pushed against the rubber blocks in the recording chamber so that the slice is not tilted (ganglion cells and photoreceptors should be in focus at the same time)</li>
</ol>
Placing the tissue in the recording rig
<ol>
<li>The recording slide is placed under the microscope and perfused with Ringer&rsquo;s solutions through a gravity perfusion syste</li>
<li>The preparation is visualized with an upright microscope with a 10x and 40x water immersion objective. </li>
<li>Again, we only use IR light to prevent bleaching of the photoreceptors. An IR camera is attached to the side port of the microscope, which is connected to a TV screen.</li>
</ol>
Visual Stimulation
Visual stimuli are programmed in Matlab using the Psychtoolbox and are projected onto the retina through the top port of the microscope using a microscopy image injector (Mbf bioscience). A Bits++ Digital Video Processor (Cambridge Research Systems) is used to obtain a 14bit luminance scale and to synchronize stimulus presentation with data acquisition. The visual stimuli used in this video are bright or dark bars (width = 460&micro;m) of 100% contrast, which were presented for two seconds on a steady uniform background (luminance = 8 * 104 photons/&micro;m/s, size: 1.84 x 1.38mm).
Electrophysiology
<ul>
<li>Patch electrodes are pulled from borosilicate glass (OD: 1.5mm, ID. .84mm) with a micropipette puller. For ganglion cells, resistances are between 2 and 4 MOhms. For bipolar cells, higher resistances between 4 and 8 MOhms are used. </li>
<li>Voltage Clamp recordings are performed using a Multiclamp 700A (Molecular Devices), amplifier and lab internal software programmed in Labview. </li>
<li>Drugs are perfused through an 8-channel perfusion system. The microperfusion electrode is placed close to the slice (the tissue should visibly move when you turn the perfusion on and off, but one needs to be careful that the pressure is not too strong so that the slice could become detached from the filter paper). During experiments the microperfusion should be constantly turned on (perfusing control solution if no drugs are needed) so that the recording does not get disturbed by switching the perfusion on and off. </li>
</ul>

<ol>
	<li>Cook, P. B., Lukasiewicz, P. D., McReynnolds, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=GABA(C)+receptors+control+adaptive+changes+in+a+glycinergic+inhibitory+pathway+in+salamander+retina.">GABA(C) receptors control adaptive changes in a glycinergic inhibitory pathway in salamander retina.</a> Journal of Neuroscience. 20, 806-812 (2000).</li><li>Heflin, S. J., Cook, P. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Narrow+and+wide+field+amacrine+cells+fire+action+potentials+in+response+to+depolarization+and+light+stimulation.">Narrow and wide field amacrine cells fire action potentials in response to depolarization and light stimulation.</a> Visual Neuroscience. 24, 197-206 (2007).</li><li>Ichinose, T., Shields, C. R., Lukasiewicz, P. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sodium+Channels+in+the+Transient+Retinal+Bipolar+Cells+Enhance+Visual+Responses+in+Ganglion+Cells.">Sodium Channels in the Transient Retinal Bipolar Cells Enhance Visual Responses in Ganglion Cells.</a> J Neurosci.. 25, 1856-1865 (2005).</li></ol>

Benefits:
<ol>
<li>All cell types are accessible </li>
<li>Easy identification of cell types, in particular if a fluorescent dye is added to the electrode solution </li>
<li>Pharmacological agents can easily reach target cells </li>
<li>The patch clamp technique allows to investigate the role of different ion channels in retinal computations. The same information can not be obtained through extracelluar spike recordings or recordings with sharp electrodes. </li>
<li>This technique can b...

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<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Multiclamp 700A</td>
<td>Patch Clamp Amplifier</td>
<td>Molecular Devices</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>SZX9</td>
<td>Dissection Micropscope</td>
<td>Olympus</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>BX50WI</td>
<td>Upright Microscope</td>
<td>Olympus</td>
<td>&nbsp;</td>
<td>Equipped with 40x, 10x water immersion objective, fluorescent filters and mercury lamp</td>
</tr>
<tr>
<td>P-97</td>
<td>Micropipette Puller</td>
<td>Sutter Instruments</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Lucivid</td>
<td>Microscope image injector</td>
<td>Mbf bioscience</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>8-channel perfusion system</td>
<td>Microperfusion</td>
<td>Parker Hannfin corporation</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Bits++</td>
<td>Digital Video Processor</td>
<td>Cambridge Research Systems</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Infrared Oculars</td>
<td>Other</td>
<td>ITT</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Adhesive silicone wells</td>
<td>Other</td>
<td>Molecular Probes</td>
<td>&nbsp;</td>
<td>20mm diameter, 1.0mm deep</td>
</tr>
<tr>
<td>Membrane Filters</td>
<td>Filter</td>
<td>Milipore</td>
<td>HAWPO1300</td>
<td>.45&#181;m HA</td>
</tr>
<tr>
<td>Borosilicate Glass Capillaries</td>
<td>Electrode glass</td>
<td>World Precision Instruments</td>
<td>1B150-4</td>
<td>&nbsp;</td>
<tr>
<td>Syringe Filters</td>
<td>Filter</td>
<td>Whatman</td>
<td>6789-0402</td>
<td>4mm filters, .2um Nylon Membrane, Polypropylene housing</td>
</tr>
<tr>
<td>IR camera</td>
<td>Micropscope mounted camera</td>
<td>Sony</td>
<td>SPT-M324</td>
<td>Extrawave HAD, B7W video camera</td>
</tr>
<tr>
<td>Picrotoxin</td>
<td>Reagent</td>
<td>Sigma</td>
<td>P1675</td>
<td>&nbsp;</td>
<tr>
<td>Strychnine</td>
<td>Reagent</td>
<td>Sigma</td>
<td>S0532</td>
<td>&nbsp;</td>
<tr>
<td>Imidazole-4-acetic acid sodium salt</td>
<td>Reagent</td>
<td>Sigma</td>
<td>I7013</td>
<td>&nbsp;</td>
<tr>
<td>Larval tiger salamanders</td>
<td>Animal</td>
<td>Charles E. Sullivan (Nashville, TN)</td>
<td>&nbsp;</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

whole-cell recordings of light evoked excitatory synaptic currents in the retinal slice

We use the whole-cell patch clamp technique to study the synaptic circuitry that underlies visual information processing in the retina. In this video, we will guide you through the process of performing whole-cell recordings of light evoked currents of individual cells in the retinal slice preparation. We use the aquatic tiger salamander as an animal model. We begin by describing the dissection of the eye and show how slices are mounted for electrophysiological recordings. Once the slice is placed in the recording chamber, we demonstrate how to perform whole-cell voltage clamp recordings. We then project visual stimuli onto the photoreceptors in the slice to elicit light-evoked current responses. During the recording we perfuse the slice with pharmacological agents, whereby an 8-channel perfusion system allows us to quickly switch between different agents.  The retinal slice preparation is widely used for patch clamp recordings in the retina, in particular to study amacrine or bipolar cells, which are not accessible in a whole-mount preparation.

Watch this Scientific Journal Video about Whole-cell Recordings of Light Evoked Excitatory Synaptic Currents in the Retinal Slice at JoVE.com

Whole-cell Recordings of Light Evoked Excitatory Synaptic Currents in the Retinal Slice

This video shows the process of whole-cell voltage clamp recordings in the retinal slice of the aquatic tiger salamander. We demonstrate the preparation of the slice as well as how to perform patch clamp recordings during visual stimulation of the retina.

We use the whole-cell patch clamp technique to study the synaptic circuitry that underlies visual information processing in the retina. In this video, we ...

whole-cell-recordings-light-evoked-excitatory-synaptic-currents

Research

JoVE Journal

Biology

12.7K Views.  Boston University. This video shows the process of whole-cell voltage clamp recordings in the retinal slice of the aquatic tiger salamander. We demonstrate the preparation of the slice as well as how to perform patch clamp recordings during visual stimulation of the retina. 

Video: Whole-cell Recordings of Light Evoked Excitatory Synaptic Currents in the Retinal Slice

Horizontal Slice Preparation of the Retina

Traditionally the vertical slice and the whole-mount preparation of the retina have been used to study the function of retinal circuits. However, many of retinal neurons, such as amacrine cells, expand their dendrites horizontally, so that the morphology of the cells is supposed to be severely damaged in the vertical slices. In the whole-mount preparation, especially for patch-clamp recordings, retinal neurons in the middle layer are not easily accessible due to the extensive coverage of glial cell (Mueller cell) s endfeets. Here, we describe the novel slicing method to preserve the dendritic morphology of retinal neurons intact. The slice was made horizontally at the inner layer of the retina using a vibratome slicer after the retina was embedded in the low-temperature melting agarose gel. In this horizontal slice preparation of the retina, we studied the function of retinal neurons compared with their morphology, by using patch-clamp recording, calcium imaging technique, immunocytochemistry, and single-cell RT-PCR.

Traditionally the vertical slice and the whole-mount preparation of the retina have been used to study the function of retinal circuits. Here, we describe the novel slicing method to preserve the dendritic morphology of retinal neurons intact.

Traditionally the vertical slice and the whole-mount preparation of the retina have been used to study the function of retinal circuits. However, many of ...

Whole Cell Recordings from Brain of Adult Drosophila

In this video, we demonstrate the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons. We begin by describing the dissecting solution and capture of the adult females used in our studies. The procedure for removing the whole brain intact, including both optic lobes, is illustrated. Dissection of the overlying trachea is also shown.  The isolated brain is not only small but needs special care in handling at this stage to prevent damage to the neurons, many of which are close to the outer surface of the tissue. We show how a special holder we developed is used to stabilize the brain in the recording chamber. A standard electrophysiology set up is used for recording from single neurons or pairs of neurons. A fluorescent image, viewed through the recording microscope, from a GAL4 line driving GFP expression (GH146) illustrates how projection neurons (PNs) are identified in the live brain. A high power Nomarski image shows a view of a single neuron that is being targeted for whole cell recording. When the brain is successfully removed without damage, the majority of the neurons are spontaneously active, firing action potentials and/or exhibiting spontaneous synaptic input. This in situ preparation, in which whole cell recording of identified neurons in the whole brain can be combined with genetic and pharmacological manipulations, is a useful model for exploring cellular physiology and plasticity in the adult CNS.

This video demonstrates the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons using standard whole cell technology. It includes images of GFP labeled cells and neurons viewed during recording.

In this video, we demonstrate the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons. We begin by ...

Major Components of the Light Microscope

The light microscope is a basic tool for the cell biologist, who should have a thorough understanding of how it works, how it should be aligned for different applications, and how it should be maintained as required to obtain maximum image-forming capacity and resolution. The components of the microscope are described in detail here.

The light microscope is a basic tool for the cell biologist, who should have a thorough understanding of how it works, how it should be aligned for different applications, and how it should be maintained as required to obtain maximum image-forming capacity and resolution. The components of the microscope are described in detail here.

The light microscope is a basic tool for the cell biologist, who should have a thorough understanding of how it works, how it should be aligned for ...

Single-cell Suction Recordings from Mouse Cone Photoreceptors

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment are open and allow cations to flow steadily inwards across the membrane, depolarizing the cell. Light exposure triggers the closure of the CNG channels, blocks the inward cation current flow, and thus results in cell hyperpolarization. Based on the polarity of photoreceptors, a suction recording method was developed in 1970s that, unlike the classic patch-clamp technique, does not require penetrating the plasma membrane 1. Drawing the outer segment into a tightly-fitting glass pipette filled with extracellular solution allows recording the current changes in individual cells upon test-flash exposure. However, this well-established "outer-segment-in (OS-in)" suction recording is not suitable for mouse cone recordings, because of the low percentage of cones in the mouse retina (3%) and the difficulties in identifying the cone outer segments. Recently, an inner-segment-in (IS-in) recording configuration was developed to draw the inner segment/nuclear region of the photoreceptor into the recording pipette 2,3. In this video, we will show how to record from individual mouse cone photoresponses using single-cell suction electrode.

We will show how to record flash responses from single mouse cones using a suction electrode.

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment ...

Whole-Cell Recording of Calcium Release-Activated Calcium (CRAC) Currents in Human T Lymphocytes

In T lymphocytes, depletion of Ca2+ from the intracellular Ca2+ store leads to activation of plasmalemmal Ca2+ channels, called Calcium Release-Activated Calcium (CRAC) channels. CRAC channels play important role in regulation of T cell proliferation and gene expression. Abnormal CRAC channel function in T cells has been linked to severe combined immunodeficiency and autoimmune diseases 1, 2 . Studying CRAC channel function in human T cells may uncover new molecular mechanisms regulating normal immune responses and unravel the causes of related human diseases. Electrophysiological recordings of membrane currents provide the most accurate assessment of functional channel properties and their regulation. Electrophysiological assessment of CRAC channel currents in Jurkat T cells, a human leukemia T cell line, was first performed more than 20 years ago 3, however, CRAC current measurements in normal human T cells remains a challenging task. The difficulties in recording CRAC channel currents in normal T cells are compounded by the fact that blood-derived T lymphocytes are much smaller in size than Jurkat T cells and, therefore, the endogenous whole-cell CRAC currents are very low in amplitude. Here, we give a step-by-step procedure that we routinely use to record the Ca2+ or Na+ currents via CRAC channels in resting human T cells isolated from the peripheral blood of healthy volunteers. The method described here was adopted from the procedures used for recording the CRAC currents in Jurkat T cells and activated human T cells 4-8.

Whole-Cell Recording of Calcium Release-Activated Calcium CRAC Currents in Human T Lymphocytes

We provide a step-by-step protocol for whole-cell patch clamp recording of Calcium Release-Activated Calcium (CRAC) currents in peripheral blood mononuclear cell-derived human T lymphocytes.

In T lymphocytes, depletion of Ca2+ from the intracellular Ca2+ store leads to activation of plasmalemmal Ca2+ channels, called Calcium Release-Activated ...

Studying Proteolysis of Cyclin B at the Single Cell Level in Whole Cell Populations

Equal distribution of chromosomes between the two daughter cells during cell division is a prerequisite for guaranteeing genetic stability 1. Inaccuracies during chromosome separation are a hallmark of malignancy and associated with progressive disease 2-4. The spindle assembly checkpoint (SAC) is a mitotic surveillance mechanism that holds back cells at metaphase until every single chromosome has established a stable bipolar attachment to the mitotic spindle1. The SAC exerts its function by interference with the activating APC/C subunit Cdc20 to block proteolysis of securin and cyclin B and thus chromosome separation and mitotic exit. Improper attachment of chromosomes prevents silencing of SAC signaling and causes continued inhibition of APC/CCdc20 until the problem is solved to avoid chromosome missegregation, aneuploidy and malignant growths1.
Most studies that addressed the influence of improper chromosomal attachment on APC/C-dependent proteolysis took advantage of spindle disruption using depolymerizing or microtubule-stabilizing drugs to interfere with chromosomal attachment to microtubules. Since interference with microtubule kinetics can affect the transport and localization of critical regulators, these procedures bear a risk of inducing artificial effects 5.
To study how the SAC interferes with APC/C-dependent proteolysis of cyclin B during mitosis in unperturbed cell populations, we established a histone H2-GFP-based system which allowed the simultaneous monitoring of metaphase alignment of mitotic chromosomes and proteolysis of cyclin B 6.
To depict proteolytic profiles, we generated a chimeric cyclin B reporter molecule with a C-terminal SNAP moiety 6 (Figure 1). In a self-labeling reaction, the SNAP-moiety is able to form covalent bonds with alkylguanine-carriers (SNAP substrate) 7,8 (Figure 1). SNAP substrate molecules are readily available and carry a broad spectrum of different fluorochromes. Chimeric cyclin B-SNAP molecules become labeled upon addition of the membrane-permeable SNAP substrate to the growth medium 7 (Figure 1). Following the labeling reaction, the cyclin B-SNAP fluorescence intensity drops in a pulse-chase reaction-like manner and fluorescence intensities reflect levels of cyclin B degradation 6 (Figure 1). Our system facilitates the monitoring of mitotic APC/C-dependent proteolysis in large numbers of cells (or several cell populations) in parallel. Thereby, the system may be a valuable tool to identify agents/small molecules that are able to interfere with proteolytic activity at the metaphase to anaphase transition. Moreover, as synthesis of cyclin B during mitosis has recently been suggested as an important mechanism in fostering a mitotic block in mice and humans by keeping cyclin B expression levels stable 9,10, this system enabled us to analyze cyclin B proteolysis as one element of a balanced equilibrium 6.

Metaphase to anaphase transition is triggered through anaphase-promoting complex (APC/C)-dependent ubiquitination and subsequent destruction of cyclin B. Here, we established a system which, following pulse-chase labeling, allows monitoring cyclin B proteolysis in entire cell populations and facilitates the detection of interference by the mitotic checkpoint.

Equal distribution of chromosomes between the two daughter cells during cell division is a prerequisite for guaranteeing genetic stability 1. Inaccuracies ...

The Slice Culture Method for Following Development of Tooth Germs In Explant Culture

Explant culture allows manipulation of developing organs at specific time points&#160;and is therefore an important method for the developmental biologist. For many organs it is difficult to access developing tissue to allow monitoring during ex vivo culture. The slice culture method allows access to tissue so that morphogenetic movements can be followed and specific cell populations can be targeted for manipulation or lineage tracing.
In this paper we describe a method of slice culture that has been very successful for culture of tooth germs in a range of species. The method provides excellent access to the tooth germs, which develop at a similar rate to that observed in vivo, surrounded by the other jaw tissues. This allows tissue interactions between the tooth and surrounding tissue to be monitored. Although this paper concentrates on tooth germs, the same protocol can be applied to follow development of a number of other organs, such as salivary glands, Meckel&#39;s cartilage, nasal glands, tongue, and ear.

Here we detail a method to culture tooth germs in mandible slices using a tissue chopper. This method allows unique access to the tooth during development, providing excellent opportunity for manipulation and lineage tracing, not available using more traditional culture methods.

Explant culture allows manipulation of developing organs at specific time points&#160;and is therefore an important method for the developmental ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

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) ...

Whole Cell Electrophysiology of Primary Cultured Murine Enterochromaffin Cells

Enterochromaffin (EC) cells in the gastrointestinal (GI) epithelium constitute the largest subpopulation of enteroendocrine cells. As specialized sensory cells, EC cells sense luminal stimuli and convert them into serotonin (5-hyroxytryptamine, 5-HT) release events. However, the electrophysiology of these cells is poorly understood because they are difficult to culture and to identify. The method presented in this paper outlines primary EC cell cultures optimized for single cell electrophysiology. This protocol utilizes a transgenic cyan fluorescent protein (CFP) reporter to identify mouse EC cells in mixed primary cultures, advancing the approach to obtaining high-quality recordings of whole cell electrophysiology in voltage- and current-clamp modes.

Enterochromaffin (EC) cells comprise a small subset of gastrointestinal epithelial cells. EC cells are electrically excitable and release serotonin, yet difficulties in culturing and identifying EC cells have limited physiological studies. The method presented here establishes a primary culture model amenable to examination of single EC cells by electrophysiology.

Enterochromaffin (EC) cells in the gastrointestinal (GI) epithelium constitute the largest subpopulation of enteroendocrine cells. As specialized sensory ...