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

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.