Name: modified roller tube method for precisely localized and repetitive intermittent imaging during long-term culture of brain slices in an enclosed system
Uploaded: 2017-12-28T13:00:00
Description: Watch this Scientific Journal Video about Modified Roller Tube Method for Precisely Localized and Repetitive Intermittent Imaging During Long-term Culture of Brain Slices in an Enclosed System at JoVE

Animal use follows approved breeding and animal use protocols that conform to the Animal Care and Use Guidelines of Colorado State University.
NOTE: The protocol below describes the preparation and culture method for the long-term incubation and intermittent imaging of hippocampal slices. A single hippocampal slice is attached to a specially prepared photoetched coverslip using a plasma clot, and then the coverslips are sealed onto the flat side of a drilled-out roller tube, which is maintaine...

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The roller tube method described here allows for long-term culturing and high-resolution live imaging of sliced brain tissue. One major issue with the slice technique as applied here is in the mounting and maintenance of slices. Coverslip coatings that support slice adhesion, promote slice thinning by enhancing the outgrowth of neurites and migration of cells out of the slice; thus, we avoided the use of these substrates. The insertion of amino groups onto the glass by treatment with 3-aminopropyltriethoxysilane improved...

Primary culture of dissociated neurons from regions of rodent brain is an important tool used by researchers to observe responses to pathologically implicated stimuli. However, such studies have the disadvantage of looking at neurons in only 2D and without their glial support system. Furthermore, unless grown under conditions of very high density (640 neurons/mm2 or about 16% of surface area) in which it becomes impossible to follow the random outgrowth of a dendrite or axon for more than a short distance from its cell body, hippocampal neuronal viability over 4 weeks declines significantly1, limiting the use of dissociated cultures for extended studies of age-related pathologies. The culturing of slices prepared from rodent brain is an attractive option that overcomes these limitations by maintaining an organized cell architecture and viability for weeks or months. Conditions for maintaining many different regions of rodent brain in slice culture have been described2.
Two major methods are widely used for long-term culture of brain slices: culturing on membranes at the air-liquid interface3 or culturing on coverslips in sealed tubes allowed to rotate in a roller incubator to provide aeration4. Slices cultured on membranes can be directly imaged with high-resolution fluorescence microscopy using an upright microscope and water immersion objectives5. Alternatively, slices cultured on membranes have been transferred to glass bottom dishes to achieve good resolution of dendritic spines using an inverted microscope6. However, both methods of imaging slices grown on membranes are open systems that require medium changes and often use antifungal and/or antibiotics to prevent or reduce contamination5,6. Slices on a membrane at the air-medium interface maintain excellent morphology and survival, but returning to precise locations during repetitive imaging at high magnification is extremely difficult unless the experiment is following only small groups of cells expressing a fluorescent marker. Although slices grown on membranes have been used with viral-mediated expression of transgenes5,6, biosafety protocols may require an enclosed culture system be employed for certain viral vectors that are used for expressing fluorescently tagged proteins and reporters of cell physiology. Furthermore, immersion objectives require decontamination between samples that will be followed in culture5. One major application of membrane interface cultures is combining high-resolution imaging with electrophysiology at single time points7.
The roller tube method with coverslips inside the plastic tube does not permit any electrophysiology or high-resolution imaging without removing the coverslip. Thus, this method has been most often applied to long-term studies in which post-fixation observations have been made8. Described here is a method that utilizes the roller tube culture technique but on drilled-out tubes with slices on coverslips that can be imaged repetitively for as long as the cultures are maintained. The enclosed system requires no medium change for imaging and utilizes photoetched coverslips to provide fiducial marks that allow imaging at high magnification, after days or weeks, the precise fields previously imaged.
We apply this method to examine changes in the rodent hippocampus, a major brain region involved in memory and learning. The rodent hippocampus is often studied as a model for pathological or age-related changes observed during development of cognitive impairment9, such as those that occur in AD. Our method is particularly well suited to study pathological changes that develop within a single slice over time in response to environmental changes, such as increases in A&#946; peptides, which is characteristic of AD8. One pathology associated with human and rodent AD brain is the presence of cofilin-actin aggregates and rods, the latter containing bundles of filaments in which cofilin and actin are in a 1:1 molar ratio10,11,12. Rods have been observed in fixed slices of rat hippocampus following A&#946; treatment, as well as within a live rodent brain slice expressing cofilin-GFP subjected to hypoxia8, and they may contribute to the synaptic dysfunction seen in AD and stroke. Here we use this new culturing method to observe the time course and distribution within slices of expressed exogenous chimeric fluorescent proteins introduced by different viruses. We then utilize the neuronal specific expression of a cofilin reporter construct to follow the development of cofilin rod and aggregate pathology in hippocampal slices in response to treatment with soluble A&#946; oligomers (A&#946;o).

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			<td height="40" style="height:40px;width:280px;">Bottoms from 15 cm culture dishes</td>
			<td style="width:220px;">VWR Scientific</td>
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			<td height="40" style="height:40px;">Polyethylene tubing (5/16&#34;; OD, 3/16&#34;; ID)</td>
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			<td>Cut to 1.8&#34; length</td>
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			<td>ACE Hardware</td>
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			<td>Bellco Glass, Inc.</td>
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			<td>A3648</td>
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		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">#5 Dumont Forceps</td>
			<td>Fine Science Tools</td>
			<td>11251-30</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">McIlwain Tissue Chopper</td>
			<td>Ted Pella, Inc.</td>
			<td>10180</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Double Edge Razor Blades</td>
			<td>Ted Pella, Inc.</td>
			<td>121-6</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Whatman Filter Paper</td>
			<td>VWR</td>
			<td>28450-182</td>
			<td>Cut into 5.8 cm diameter circles</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Poly-chloro-trifluoro-ethylene (Aclar)</td>
			<td>Ted Pella, Inc.</td>
			<td>10501-10</td>
			<td>Cut into 5.8 cm diameter circles</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">#21 Surgical Blade</td>
			<td>VWR Scientific</td>
			<td>25860-144</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">#5 Dumont Forceps</td>
			<td>Fine Science Tools</td>
			<td>11251-30</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Spatula, stainless with tapered end</td>
			<td>VWR</td>
			<td>82027-518</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Gey&#39;s Balanced Salt Solution</td>
			<td>Sigma-Aldrich</td>
			<td>G9779&#160;</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">&#160;Glucose</td>
			<td>ThermoFisher Scientific</td>
			<td>15023-021</td>
			<td>25% (w/v) Solution, 0.2 mm filter sterilized</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Chicken Plasma</td>
			<td>Cocalico Biologicals</td>
			<td>30-0300-5L</td>
			<td>Rehydrate in sterile water, centrifuge at 2500 x g 30 min at 4 &#176;C, quick freeze aliquots in liquid nitrogen and store at&#160; -80 &#176;C.</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Thrombin, Bovine</td>
			<td>Fisher</td>
			<td>60-516-01KU</td>
			<td>150 units /ml in GBSS/Glucose, quick freeze aliquots in liquid nitrogen and store at -80 &#176;C.</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Cell Roller System</td>
			<td>Bellco Biotech</td>
			<td>SciERA</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Roller Incubator</td>
			<td>Forma</td>
			<td>Model 3956</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">N21-MAX</td>
			<td>ThermoFisher Scientific</td>
			<td>AR008</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Pen/Strep (100X)</td>
			<td>ThermoFisher Scientific</td>
			<td>15140122</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">200 mM Glutamine</td>
			<td>ThermoFisher Scientific</td>
			<td>25030081</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Glucose</td>
			<td>ThermoFisher Scientific</td>
			<td>15023-021</td>
			<td>25% (w/v) Solution, 0.2 mm filter sterilized</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Neurobasal A</td>
			<td>ThermoFisher Scientific</td>
			<td>10888-022&#160;</td>
			<td>Complete Medium: 48 mL Neurobasal A, 1 mL N21-MAX, 0.625 mL 200 mM Glutamine, 0.180 mL 25% Glucose, 0.250 mL 100x pen/strep.</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Third generation lentivirus packaging</td>
			<td>Life Technologies</td>
			<td>K4975-00</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">159 K cutoff centrifugal filters (Centricon)</td>
			<td>EMD Millipore</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Lentiviral cloning system (InFusion)</td>
			<td>Clonetech</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Plasmids 30323, 50856, 51279</td>
			<td>Addgene</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Neuronal cell viability dye (NeuO)</td>
			<td>Stemcell technologies</td>
			<td>1801</td>
			<td>Thaw once and quick freeze in 4 &#181;L aliquots. Store at -20 &#176;C</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Inverted microscope</td>
			<td>Olympus</td>
			<td>IX83</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Microscope objectives</td>
			<td>Olympus</td>
			<td>air: 4X, 20; oil: 40X, 60X,</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Spinning disc confocal system</td>
			<td>Yokagawa</td>
			<td>CSU22</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Microscope EMCCD camera</td>
			<td>Photometrics</td>
			<td>Cascade II</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Linear encoded (x,y), piezo z flat top stage</td>
			<td>ASI</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Microscope lasers and integration</td>
			<td>Intelligent Imaging Innovations</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">HEK293T cells</td>
			<td>American Type Culture Collection</td>
			<td>CRL-3216</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Human Plasmin</td>
			<td>Sigma Aldrich</td>
			<td>P1867</td>
			<td>0.002 U/mL in 0.1% bovine serum albumin (0.2 mm filter sterilized), quick freeze in liquid nitrogen and store at -80 &#176;C.</td>
		</tr>
	</tbody>
</table>

modified roller tube method for precisely localized and repetitive intermittent imaging during long-term culture of brain slices in an enclosed system

Cultured rodent brain slices are useful for studying the cellular and molecular behavior of neurons and glia in an environment that maintains many of their normal in vivo interactions. Slices obtained from a variety of transgenic mouse lines or use of viral vectors for expression of fluorescently tagged proteins or reporters in wild type brain slices allow for high-resolution imaging by fluorescence microscopy. Although several methods have been developed for imaging brain slices, combining slice culture with the ability to perform repetitive high-resolution imaging of specific cells in live slices over long time periods has posed problems. This is especially true when viral vectors are used for expression of exogenous proteins since this is best done in a closed system to protect users and prevent cross contamination. Simple modifications made to the roller tube brain slice culture method that allow for repetitive high-resolution imaging of slices over many weeks in an enclosed system are reported. Culturing slices on photoetched coverslips permits the use of fiducial marks to rapidly and precisely reposition the stage to image the identical field over time before and after different treatments. Examples are shown for the use of this method combined with specific neuronal staining and expression to observe changes in hippocampal slice architecture, viral-mediated neuronal expression of fluorescent proteins, and the development of cofilin pathology, which was previously observed in the hippocampus of Alzheimer's disease (AD) in response to slice treatment with oligomers of amyloid-&#946; (A&#946;) peptide.

To determine how accurately fiducial marks can be utilized to reimage the same cells within the same fields over time, we examined slices grown on photoetched coverslips (Figure 6A). Neurons were visualized by staining with a vital dye (100 nM for 2 h; does not stain non-neuronal cells), which disappears from neurons over time without harming the cells25. We identified a fiducial mark in a single grid square (<strong c...

Watch this Scientific Journal Video about Modified Roller Tube Method for Precisely Localized and Repetitive Intermittent Imaging During Long-term Culture of Brain Slices in an Enclosed System at JoVE

Modified Roller Tube Method for Precisely Localized and Repetitive Intermittent Imaging During Long-term Culture of Brain Slices in an Enclosed System

Presented here is a modified roller tube method for culturing and intermittent high-resolution imaging of rodent brain slices over many weeks with precise repositioning on photoetched coverslips. Neuronal viability and slice morphology are well maintained. Applications of this fully enclosed system using viruses for cell-type specific expression are provided.

Cultured rodent brain slices are useful for studying the cellular and molecular behavior of neurons and glia in an environment that maintains many of ...

The overall goal of this modified roller tube method for brain slice culture is to provide an enclosed culture system for long-term slice survival. with the capability for repetitive high-resolution florescence imaging. This method can help answer key questions in neuronal development and neurodegenerative disorders that require observations of the same neurons over time, such as localizing key proteins involved in synapse formation or loss.The main advantages of this technique are the fully enclosed culture system that allows safe use of viruses for protein expression and the use of photo etched coverslips to localize the same cells for imaging. We first had the idea for this method when we were culturing slices on coverslips inside roller tubes, but needed a better way to visualize changes occurring in living slices over time. The widespread use of viral vectors to express transgenes and specific cell populations for microscope imaging drove us to develop an enclosed system to protect users and eliminate contamination of microscope objectives.The ability to follow the identical population of cells within brain slices over time allows for studies on development of pathological changes and synapse alteration in rodent models of human neurological disorders. To begin prepare the roller tube rack and make a jig to assist with drilling a hole into the roller tubes as described in the accompanying text protocol. Using the jig, hold in position a flat sided, one centimeter, plastic culture tube, so that the flat side is facing up.Then drill a six millimeter diameter hole with the center one centimeter from the bottom and centered between the sides of the tube. With the swiveling deburring tool smooth the edges of the hole and make four grooves on the inside edge of the hole to facilitate draining of the hole during rotation. Next, rinse the tubes with 70%ethanol and then air-dry them in a biological safety cabinet.Sterilize the tubes in some 12 millimeter punched adhesive silicone rubber disks for 40 minutes under the UV lamp. After 20 minutes reposition the tubes and disks so that all of the exposed surfaces are sterilized. Next peel off the white backing from the adhesive disk, align the holes and affix the silicone rubber to the outside of the tube.Once the slices have been obtained place two microliters of chicken plasma on the center of the photo etched side of a prepared coverslip. Spread the plasma slightly to achieve a three to four milliliter diameter spot. Then use a sterile, narrow-tipped spatula to lift a prepared brain slice.Use closed forceps to keep the slice on the spatula tip while lifting the slice. Touch the spatula to the plasma spot on the coverslip and with closed forceps push the slice onto the coverslip. Add a thin layer of chick plasma on the coverslip before placing the slice and a minimal amount of thrombin treated plasma to cover to cover the slice.If the slice floats it will not remain adhered when the cloud is removed. Next mix 2.5 microliters of plasma and 2.5 microliters of thrombin in a separate tube. Quickly place 2.5 microliters of this mixture over and around the slice and pipette up and down gently to mix it.Remove the clear, plastic covering from the exposed side of the silicone rubber adhesive previously affixed to the roller tube. Then place the coverslip with the brain slice onto the adhesive aligning the slice within the hole. To ensure adhesion apply soft even pressure to the coverslip with the thumb, by pressing the coverslip down evenly, and holding it for a about one minute while transferring it to the biological safety cabinet.The pressure on the coverslip to adhere it to the tube without leaking must be just right and maintained for long enough to eliminate air channels in the salient. Too much pressure will crack the coverslip. In a biological safety cabinet, add 0.8 milliliters of complete neurobasal a culture medium to each tube.Then flow a a 5%carbon dioxide 95%air mixture through a sterile cotton plugged Pasteur pipette held securely by a clamp. Use this to flush the roller tube with the gas mixture and rapidly cap the tube as it is withdrawn from around the pipette. Next label the tubes with the slice number and rack number.Insert the tubes into a roller rack;ensuring that they are geometrically-balanced. If there is an odd number of tubes add blank tubes to balance. Place the racks in a 35 degree Celsius roller incubator, with rollers turning the roller rack at 10 to 13 revolutions per hour.Tilt the incubator back approximately 5 degrees by lifting its front on to a board. Transfer a tube with a slice culture to a custom-made tube holder like the one shown here. Then the place the tube holder into the stage adapter to keep the coverslip perpendicular to the objective during imaging.Next push the slider on the stage adapter tight against the tube to hold the tube in position. Using a four times objective under bright field trans-illumination, focus on the photo etched grid period under the slice. For the initial imaging session, quickly scan around the slice to locate and record the number for the various regions where higher magnification imaging is desired.Once the regions of interest have been identified, move the stage to the first grid square containing a region of interest. Then switch to the 20 strength error objective and locate a fiducial mark. Next switch to a higher power objective, localize the same fiducial mark, and record the x and y position of the stage.Move the stage to find the other fields of interest nearby and record their x and y offsets from the fiducial mark. The offset positions allow for consistent pinpoint of the same coverslip location when the slice is imaged, even though the original x, y setting of the fiducial mark changes when the tube or stage adapter are removed and replaced. Capture an image Z-stick within each selected field using the either the microscope objective control or a Piezo stage control if one is available.Using a neuronal vital dye, the confocal stack of images show here, highlights the 3D architecture of both neurites and the cell body, but the stain is excluded from the nucleus. To examine time-dependent changes in growth slice morphology and viability within slices during long-term culture, the same slice was labeled with fluorescent neuronal vital dye once per week, 24 hours before imaging. To demonstrate the reproducibility of imaging the identical cells over time, the same field of a slice labeled with neuronal vital dye on day 13, was imaged on days 14, 15, 16, and 17.The position of three cells on each day is shown by a symbol over the nucleus. In addition to the neuronal vital dye, viral-mediated fluorescent proteins can be utilized with this system to label cells. Here both a Calcium sensitive reporter and Cofilin-containing rods are able to be clearly identified 10 days post-infection.Successful implementation of this procedure requires preparation in advance of all tubes, coverslips, and solutions. Being able to obtain slices in a short post-mortem interval is also crucial to success and requires much practice. Once mastered this technique can be done in about 90 minutes to obtain 12-18 hippocampal slices from a mouse pup, including times for dissection and plating.Following this procedure, brain slices can be infected at various times with viruses for achieving cell-type specific expression of fluorescently tagged proteins or silence for gene expression with small interfering RNA or short hairpin RNA to probe gene function. Don't forget that working with recombinant viruses can be hazardous and precautions such as wearing eye protection should always be taken when working with these reagents.

modified-roller-tube-method-for-precisely-localized-repetitive

Research

JoVE Journal

Neuroscience

A Polished and Reinforced Thinned-skull Window for Long-term Imaging of the Mouse Brain

In vivo imaging of cortical function requires optical access to the brain without disruption of the intracranial environment. We present a method to form a polished and reinforced thinned skull (PoRTS) window in the mouse skull that spans several millimeters in diameter and is stable for months. The skull is thinned to 10 to 15 &mu;m in thickness with a hand held drill to achieve optical clarity, and is then overlaid with cyanoacrylate glue and a cover glass to: 1) provide rigidity, 2) inhibit bone regrowth and 3) reduce light scattering from irregularities on the bone surface. Since the skull is not breached, any inflammation that could affect the process being studied is greatly reduced. Imaging depths of up to 250 &mu;m below the cortical surface can be achieved using two-photon laser scanning microscopy. This window is well suited to study cerebral blood flow and cellular function in both anesthetized and awake preparations. It further offers the opportunity to manipulate cell activity using optogenetics or to disrupt blood flow in targeted vessels by irradiation of circulating photosensitizers.

We present a method to form an imaging window in the mouse skull that spans millimeters and is stable for months without inflammation of the brain. This method is well suited for longitudinal studies of blood flow, cellular dynamics, and cell/vascular structure using two-photon microscopy.

In vivo imaging of cortical function requires optical access to the brain without disruption of the intracranial environment. We present a method to form ...

Improved Preparation and Preservation of Hippocampal Mouse Slices for a Very Stable and Reproducible Recording of Long-term Potentiation

Long-term potentiation (LTP) is a type of synaptic plasticity characterized by an increase in synaptic strength and believed to be involved in memory encoding. LTP elicited in the CA1 region of acute hippocampal slices has been extensively studied. However the molecular mechanisms underlying the maintenance phase of this phenomenon are still poorly understood. This could be partly due to the various experimental conditions used by different laboratories. Indeed, the maintenance phase of LTP is strongly dependent on external parameters like oxygenation, temperature and humidity. It is also dependent on internal parameters like orientation of the slicing plane and slice viability after dissection.
The optimization of all these parameters enables the induction of a very reproducible and very stable long-term potentiation. This methodology offers the possibility to further explore the molecular mechanisms involved in the stable increase in synaptic strength in hippocampal slices. It also highlights the importance of experimental conditions in in vitro investigation of neurophysiological phenomena.

This paper presents a complete methodology to prepare and preserve in vitro acute hippocampal slices from adult mice. This protocol allows the recording of very stable long-lasting long-term potentiation (LTP) for more than 8 hours with a success rate of 95%.

Long-term potentiation (LTP) is a type of synaptic plasticity characterized by an increase in synaptic strength and believed to be involved in memory ...

In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

The subventricular zone (SVZ) is one of the main neurogenic niches in the postnatal brain. Here, neural progenitors proliferate and give rise to neuroblasts able to move along the rostral migratory stream (RMS) towards the olfactory bulb (OB). This long-distance migration is required for the subsequent maturation of newborn neurons in the OB, but the molecular mechanisms regulating this process are still unclear. Investigating the signaling pathways controlling neuroblast motility may not only help understand a fundamental step in neurogenesis, but also have therapeutic regenerative potential, given the ability of these neuroblasts to target brain sites affected by injury, stroke, or degeneration.
In this manuscript we describe a detailed protocol for in vivo postnatal electroporation and subsequent time-lapse imaging of neuroblast migration in the mouse RMS. Postnatal electroporation can efficiently transfect SVZ progenitor cells, which in turn generate neuroblasts migrating along the RMS. Using confocal spinning disk time-lapse microscopy on acute brain slice cultures, neuroblast migration can be monitored in an environment closely resembling the in vivo condition. Moreover, neuroblast motility can be tracked and quantitatively analyzed. As an example, we describe how to use in vivo postnatal electroporation of a GFP-expressing plasmid to label and visualize neuroblasts migrating along the RMS. Electroporation of shRNA or CRE recombinase-expressing plasmids in conditional knockout mice employing the LoxP system can also be used to target genes of interest. Pharmacological manipulation of acute brain slice cultures can be performed to investigate the role of different signaling molecules in neuroblast migration. By coupling in vivo electroporation with time-lapse imaging, we hope to understand the molecular mechanisms controlling neuroblast motility and contribute to the development of novel approaches to promote brain repair.

In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

Neuroblast migration is a fundamental event in postnatal neurogenesis. We describe a protocol for efficient labeling of neuroblasts by in vivo postnatal electroporation and subsequent visualization of their migration using time-lapse imaging of acute brain slices. We include a description for the quantitative analysis of neuroblast dynamics by video tracking.

The subventricular zone (SVZ) is one of the main neurogenic niches in the postnatal brain. Here, neural progenitors proliferate and give rise to ...

Long-term Time Lapse Imaging of Mouse Cochlear Explants

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building the highly ordered, complex structure of the mammalian cochlea proceeds for around ten days. In order to study changes in gene expression over this period and their response to pharmaceutical or genetic manipulation, long-term imaging is necessary. Previously, live imaging has typically been limited by the viability of explanted tissue in a humidified chamber atop a standard microscope. Difficulty in maintaining optimal conditions for culture growth with regard to humidity and temperature has placed limits on the length of imaging experiments. A microscope integrated into a modified tissue culture incubator provides an excellent environment for long term-live imaging. In this method we demonstrate how to establish embryonic mouse cochlear explants and how to use an incubator microscope to conduct time lapse imaging using both bright field and fluorescent microscopy to examine the behavior of a typical embryonic day (E) 13 cochlear explant and Sox2, a marker of the prosensory cells of the cochlea, over 5 days.

Live imaging of the embryonic mammalian cochlea is challenging because the developmental processes at hand operate on a temporal gradient over ten days. Here we present a method for culturing and then imaging embryonic cochlear explant tissue taken from a fluorescent reporter mouse over five days.

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building ...

Regioselective Biolistic Targeting in Organotypic Brain Slices Using a Modified Gene Gun

Transfection of DNA has been invaluable for biological sciences and with recent advances to organotypic brain slice preparations, the effect of various heterologous genes could thus be investigated easily while maintaining many aspects of in vivo biology. There has been increasing interest to transfect terminally differentiated neurons for which conventional transfection methods have been fraught with difficulties such as low yields and significant losses in viability. Biolistic transfection can circumvent many of these difficulties yet only recently has this technique been modified so that it is amenable for use in mammalian tissues.
New modifications to the accelerator chamber have enhanced the gene gun&#39;s firing accuracy and increased its depths of penetration while also allowing the use of lower gas pressure (50 psi) without loss of transfection efficiency as well as permitting a focused regioselective spread of the particles to within 3 mm. In addition, this technique is straight forward and faster to perform than tedious microinjections. Both transient and stable expression are possible with nanoparticle bombardment where episomal expression can be detected within 24 hr and the cell survival was shown to be better than, or at least equal to, conventional methods. This technique has however one crucial advantage: it permits the transfection to be localized within a single restrained radius thus enabling the user to anatomically isolate the heterologous gene&#39;s effects. Here we present an in-depth protocol to prepare viable adult organotypic slices and submit them to regioselective transfection using an improved gene gun.

Recent improvements in organotypic brain slice preparations have permitted their exploitation for biotechnological applications. Organotypic slices maintain local structural characteristics of in vivo biology, including functional synaptic connections. Here we present a regioselective biolistic delivery method to label and genetically manipulate these slices.

Transfection of DNA has been invaluable for biological sciences and with recent advances to organotypic brain slice preparations, the effect of various ...

Isolation, Culture and Long-Term Maintenance of Primary Mesencephalic Dopaminergic Neurons From Embryonic Rodent Brains

Degeneration of mesencephalic dopaminergic (mesDA) neurons is the pathological hallmark of Parkinson&#8217;s diseae. Study of the biological processes involved in physiological functions and vulnerability and death of these neurons is imparative to understanding the underlying causes and unraveling the cure for this common neurodegenerative disorder. Primary cultures of mesDA neurons provide a tool for investigation of the molecular, biochemical and electrophysiological properties, in order to understand the development, long-term survival and degeneration of these neurons during the course of disease. Here we present a detailed method for the isolation, culturing and maintenance of midbrain dopaminergic neurons from E12.5 mouse (or E14.5 rat) embryos. Optimized cell culture conditions in this protocol result in presence of axonal and dendritic projections, synaptic connections and other neuronal morphological properties, which make the cultures suitable for study of the physiological, cell biological and molecular characteristics of this neuronal population.

The causes of degeneration of midbrain dopaminergic neurons during Parkinson&#8217;s disease are not fully understood. Cellular culture systems provide an essential tool for study of the neurophysiological properties of these neurons. Here we present an optimized protocol, which can be utilized for in vitro modeling of neurodegeneration.

Degeneration of mesencephalic dopaminergic (mesDA) neurons is the pathological hallmark of Parkinson&#8217;s diseae. Study of the biological processes ...

Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans

Behavior is controlled by the nervous system. Calcium imaging is a straightforward method in the transparent nematode Caenorhabditis elegans to measure the activity of neurons during various behaviors. To correlate neural activity with behavior, the animal should not be immobilized but should be able to move. Many behavioral changes occur during long time scales and require recording over many hours of behavior. This also makes it necessary to culture the worms in the presence of food. How can worms be cultured and their neural activity imaged over long time scales? Agarose Microchamber Imaging (AMI) was previously developed to culture and observe small larvae and has now been adapted to study all life stages from early L1 until the adult stage of C. elegans. AMI can be performed on various life stages of C. elegans. Long-term calcium imaging is achieved without immobilizing the animals by using short externally triggered exposures combined with an electron multiplying charge-coupled device (EMCCD) camera recording. Zooming out or scanning can scale up this method to image up to 40 worms in parallel. Thus, a method is described to image behavior and neural activity over long time scales in all life stages of C. elegans.

Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans

Imaging behavior and neural activity over long time scales without immobilization of the animal is a prerequisite to understand behavior. Agarose microfluidic chambers imaging (AMI) can be used to image neural activity and behavior for all life stages of Caenorhabditis elegans.

Behavior is controlled by the nervous system. Calcium imaging is a straightforward method in the transparent nematode Caenorhabditis elegans to measure ...

A Simplified Method for Ultra-Low Density, Long-Term Primary Hippocampal Neuron Culture

Culturing primary hippocampal neurons in vitro facilitates mechanistic interrogation of many aspects of neuronal development. Dissociated embryonic hippocampal neurons can often grow successfully on glass coverslips at high density under serum-free conditions, but low density cultures typically require a supply of trophic factors by co-culturing them with a glia feeder layer, preparation of which can be time-consuming and laborious. In addition, the presence of glia may confound interpretation of results and preclude studies on neuron-specific mechanisms. Here, a simplified method is presented for ultra-low density (~2,000 neurons/cm2), long-term (&#62;3 months) primary hippocampal neuron culture that is under serum free conditions and without glia cell support. Low density neurons are grown on poly-D-lysine coated coverslips, and flipped on high density neurons grown in a 24-well plate. Instead of using paraffin dots to create a space between the two neuronal layers, the experimenters can simply etch the plastic bottom of the well, on which the high density neurons reside, to create a microspace conducive to low density neuron growth. The co-culture can be easily maintained for &#62;3 months without significant loss of low density neurons, thus facilitating the morphological and physiological study of these neurons. To illustrate this successful culture condition, data are provided to show profuse synapse formation in low density cells after prolonged culture. This co-culture system also facilitates the survival of sparse individual neurons grown in islands of poly-D-lysine substrates and thus the formation of autaptic connections.

Low density cultures of primary hippocampal neurons usually require glia feeder layer to supply neurotrophic factors and sustain longevity. We describe here a simplified method to culture ultra-low density neurons on glass coverslips in the presence of a high density neuronal feeder layer, which facilitates investigation of specific neuronal-autonomous mechanisms.

Culturing primary hippocampal neurons in vitro facilitates mechanistic interrogation of many aspects of neuronal development. Dissociated embryonic ...

Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have validated the utility of this method for enhancing neuronal preservation and overall brain slice viability. The implementation of this technique by early adopters has facilitated detailed investigations into brain function using diverse experimental applications and spanning a wide range of animal ages, brain regions, and cell types. Steps are outlined for carrying out the protective recovery brain slice technique using an optimized NMDG artificial cerebrospinal fluid (aCSF) media formulation and enhanced procedure to reliably obtain healthy brain slices for patch clamp electrophysiology. With this updated approach, a substantial improvement is observed in the speed and reliability of gigaohm seal formation during targeted patch clamp recording experiments while maintaining excellent neuronal preservation, thereby facilitating challenging experimental applications. Representative results are provided from multi-neuron patch clamp recording experiments to assay synaptic connectivity in neocortical brain slices prepared from young adult transgenic mice and mature adult human neurosurgical specimens. Furthermore, the optimized NMDG protective recovery method of brain slicing is compatible with both juvenile and adult animals, thus resolving a limitation of the original methodology. In summary, a single media formulation and brain slicing procedure can be implemented across various species and ages to achieve excellent viability and tissue preservation.

Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

This protocol demonstrates the implementation of an optimized N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. A single media formulation is used to reliably obtain healthy brain slices from animals of any age and for diverse experimental applications.

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have ...

Recording Synaptic Plasticity in Acute Hippocampal Slices Maintained in a Small-volume Recycling-, Perfusion-, and Submersion-type Chamber System

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful analysis of synaptic transmission modulation when performing field potential or intracellular recordings. This manuscript describes methodological aspects that might be helpful in improving experimental conditions for the maintenance of acute brain slices and for recording field excitatory postsynaptic potentials in a commercially available submersion chamber with an outflow-carbogenation unit. The outflow-carbogenation helps to stabilize the oxygen level in experiments that rely on the recycling of a small buffer reservoir to enhance the cost-efficiency of drug experiments. In addition, the manuscript presents representative experiments that examine the effects of different carbogenation modes and stimulation paradigms on the activity-dependent synaptic plasticity of synaptic transmission.

This protocol describes the stabilization of the oxygen level in a small volume of recycled buffer and methodological aspects of recording activity-dependent synaptic plasticity in submerged acute hippocampal slices.

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful ...