Name: slice preparation, organotypic tissue culturing and luciferase recording of clock gene activity in the suprachiasmatic nucleus
Uploaded: 2011-02-15T18:00:00
Duration: 10 min 6 s
Description: Watch this Scientific Journal Video about Slice Preparation, Organotypic Tissue Culturing and Luciferase Recording of Clock Gene Activity in the Suprachiasmatic Nucleus at JoVE.com

This work was funded by The Swedish Medical Research Council (K2009-75SX-21028-01-3, K2008-61X-20700-01-3); FONCICYT 000000000091984; the foundations of Jeansson, S&ouml;derstr&ouml;m K&ouml;nigska sjukhemmet, M&auml;rtha Lundqvist and Sigurd och Elsa Goljes Minne; and Swedish Society of Medicine SLS-95151. Professor Gene D. Block, UCLA, is gratefully acknowledged for valuable comments on the manuscript. We thank Dr Michael And&auml;ng and Dr Helena Johard for the HCN channel blocker, and Prof. Abdel El-Manira for providing video microscope.
Ethical considerations:
All experiments on animals were performed in accordance with the guidelines and regulations set forth by Karolinska Institutet and "Stockholm's Norra Djurf&ouml;rs&ouml;ksetiska N&auml;mnd". All animal experiments are performed with the intention to minimize any possible stress or discomfort to the animal.

1. Solution Preparation
<ol>
<li>Culture medium with air-buffering capacity
<ol>
<li>Fill a 1 liter bottle with approximately 800 mL sterile H2O (autoclaved milliQ H2O).</li>
<li>While stirring, add and mix following substances: 1 container low glucose, serum-free DMEM 2902 powder without sodium bicarbonate and phenol red (phenol red interferes with the bioluminescence signal), 20 mL B27 supplement 50x, 4.7 mL of a 7.5% NaHCO3 solution (or 0.35 g NaHCO3), 10 mL HEPES 1M, 2.5 mL PenStrep 10,000 U/ mL and 3.5 g D-glucose. Let the medium stir until the ingredients are completely dissolved. 
The final medium (1 liter) will contain: 
1x DMEM, 1x B27 supplement, 4.2 mM NaHCO3, 10 mM HEPES, 25 U/ mL penicillin, 25 U/ mL streptomycin and 19 mM D-glucose.</li>
<li>Adjust pH to 7.2, using NaOH to decrease or HCl to increase the pH, and bring volume up to 1 liter with sterile H2O.</li>
<li>Adjust osmolality with H2O (decrease osmolality) or D-glucose (increase osmolality). The osmolality must be between 285-315 mOsm/Kg but the optimal range is 300-310 mOsm/Kg. Do not to dilute the medium more than 5 %.</li>
<li>Filter the culture medium using Corning sterile vaccum filtration sets (e.g. 2 x 500 mL with pore size 0.22 &mu;m) in a sterile hood. Keep the medium in 4&deg;C and protect it from light using aluminum foil. We recommend that the medium is used within 3 months.</li>
</ol>
</li>
<li>Hank's balanced salt solution (HBSS) buffer with supplements (for cutting slices)
<ol>
<li>Fill a 1 liter glass bottle with ~600 mL sterile (autoclaved milliQ) H2O and add the following substances: 100 mL HBSS 10x stock, 10 mL Penicillin-Streptomycin 10,000 U/ mL, 5 mL of a 7.5% NaHCO3 solution and 10 mL HEPES 1M. 
The final cutting solution will contain: 
1x HBSS, 10 mM HEPES, 4.5 mM NaHCO3, 100 U/ mL penicillin and 100 U/ mL streptomycin.</li>
<li>Check pH and, if necessary, adjust pH to 7.2 and bring the volume up to 1 liter with sterile H2O.</li>
<li>Check osmolality, which must be between 285-315 mOsm/Kg.</li>
<li>Cool the HBSS to 4&deg;C. The HBSS buffer needs to be very cold (4&deg;C) during the slicing/cutting procedure in order to bring down the metabolism and preserve tissue viability.</li>
</ol>
</li>
</ol>
2. Preparations Before Cutting and Culturing Slices
<ol>
<li>The tissues are cultured in a warm dry chamber without CO2. To avoid drying out the cultures need to be sealed with cover glasses attached to the dishes by grease. For this purpose, fill 5 mL syringes with silicone-based vacuum grease. Cover the syringe tips with small pieces of aluminum foil and autoclave them. Also, autoclave filter papers (used during slicing procedure).</li>
<li>Right before slicing procedure, apply autoclaved vacuum grease on the top ring surface of 35 mm petri dishes. Prepare a separate petri dish for each slice culture.</li>
<li>Before starting slicing procedure all non-sterile instruments, razor blades, vibratome blades, cover glasses and other material used for the slice and culture procedure have to be sterilized. Spray all non-sterile equipment with 70% ethanol. Expose the instruments and the greased petri dishes with UV in a sterile hood for at least 30 minutes UV exposure before culturing.</li>
<li>At the same time, fill a Falcon tube with air-buffering culture medium (count 1.2 mL for each culture; for 8 cultures prepare for instance 10 mL medium) and add freshly thawed luciferin (0.1 M stock solution; Promega, Madison, WI) (10 &mu;L luciferin to 10 mL medium; final concentration 0.1 mM).</li>
<li>The luciferin is light sensitive and both stock solutions and the luciferin-containing medium need to be protected from light. Place the Falcon tube with luciferin-medium in a dark 36-37&deg;C heating chamber. If appropriate, the luciferin can also be added directly into the culture dishes at time of cutting. If luciferin is not added, there will be no light reaction between luciferase and luciferin and therefore no signal from the tissue.</li>
<li>After UV exposure, attach a sterile blade to the vibratome.</li>
</ol>
3. SCN Slicing Procedure
The following procedure describes slicing of adult, typically 2-4 months old, C57/BL6 mice. Please note: the cutting procedure, as well as light exposure of the animal during the night, can differentially reset the phase of the SCN. To avoid this, the cutting and culturing should be performed during the light hours, preferably between ZT 6-12, when no substantial phase shifts due to the procedure occur 12. If SCN has to be sampled in darkness, 3.1 and 3.2 have to be performed in red light or with night goggles to avoid light-induced phase shifts.
<ol>
<li>Anesthetize the mouse preferably by isofluorane (Baxter) in a glass chamber. When the animal has lost its pain reflexes (check by pinching with the nails in the paw) but has not yet stopped breathing (to maintain oxygen supply as long as possible), rapidly decapitate the head with a pair of scissors or similar. 
Please note: in some countries CO2 exposure (hypercapnia) is still permitted as anesthetic method, although it has been reported to promote animal anxiety. In addition, cervical dislocation may be required before decapitation. Please follow local legislations when euthanizing animals.</li>
<li>Remove the eyes from the head with scissors in order to prevent strain and further excitation of the optic nerves, which can damage the SCN.</li>
<li>If still attached, remove the last cervical vertebrate with scissors, remove the skin (Fig 1a) and make two cuts with a fine pair of scissors (for instance iris scissors), one cut at each side of the skull along the sides, thus making a removable "lid".</li>
<li>Open the skull with a micro rongeur tool (fine dissecting scissors can also be used on a mouse) and remove all bone until the olfactory bulbs can be seen. Work upwards, never pressing down the brain with the tool, in order to prevent damage of the ventral SCN (Fig 1b).</li>
<li>Carefully cut the optic nerve between the olfactory bulbs and the hemispheres using fine micro dissecting spring scissors. Make sure that the nerve is completely cut since stretching of the optic nerve can lead to damage in the SCN and ruptured slices.</li>
<li>Turn the head upside down and let the intact brain fall out (if still attached to the brain, other cranial nerves may need to be cut caudal to the optic nerve) into a container (for instance a glass petri dish &#8776; 10 cm) filled with 50-100 mL cold HBSS allowing rapid cooling of the brain. Ideally, the two optic nerves should remain intact (Fig 1c). Keep the brain in the HBSS 30-60 seconds to make sure the brain is cooled.</li>
<li>Use a spoon or similar and place the chilled brain, dorsal surface up, on a sterile cutting surface (for example a glass petri dish lid turned upside down). In order to prepare a coronal cut, make a perpendicular cut with sterile razor blades or scalpels between the cerebral hemispheres and the cerebellum, thus removing the cerebellum.</li>
<li>Apply superglue on the dry platform belonging to the vibroslicer/vibratome.</li>
<li>Pick up the brain (cerebral hemispheres without cerebellum and without olfactory bulbs) by inserting a sharp, curved forceps in the rostral part and carefully dry off the HBSS on a sterile filter paper, still holding the brain with the forceps. (Instead of inserting a forceps, a small piece of sterile filter paper can be used to attach and transfer the brain).</li>
<li>Fix the hemispheres onto the glued platform with the rostral tip upwards and ventral surface closest to the cutting blade. Attach the platform in the holder belonging to the vibratome (for instance from Campden Instruments, UK) and immediately fill it up with cold HBSS. If the perpendicular cut is made properly, the hemispheres should stand straight up thus providing a good angle necessary for making a coronal cut containing the bilateral SCN.</li>
<li>In order to reach the target SCN area, start cutting off thicker sections (500-800 &mu;m) of the hemispheres at high or maximum speed of the vibratome. Moving the blade can be fairly quick in the beginning before reaching the hypothalamus but should be slowed down when the optic chiasm becomes visible (high frequency of the vibrating blade and slow movement horizontally decreases cell damage during slicing, thus increasing slice viability). Reduce the sections to 100 &mu;m when the optic chiasm becomes larger (wider) and the anterior commissure becomes smaller. In order to acquire a mid-SCN section, work yourself "down" (caudal direction) until the two SCN nuclei start to appear. A magnifying glass may be necessary for visualizing the nuclei. Please note: the SCN in the mouse brain is located more caudal of the optic chiasm as compared to the rat brain.</li>
<li>When the desired level of SCN has been reached (the SCN will at this point appear as more defined, round or almond shaped structures; Fig 1d; approximately Bregma -0.46- -0.70 mm for the center region of SCN in mouse 13, Bregma -0.92- -1.40 mm for rat 14), cut the SCN section (Fig 1e). Suitable thickness of the slice is for mouse 250 &plusmn; 50 &mu;m; for rat 350 &plusmn; 50 &mu;m.</li>
<li>Using a soft brush, lift and transfer the SCN slice to a lid from a medium sized petri dish filled with cold HBSS, placed under a dissection microscope or stereoscope. Check under magnification if the bilateral SCN is clearly visible. If the mid-region of the SCN will be chosen (which may not necessarily be the only "optimal" slice level but that we consider is the easiest way to standardize the slicing procedure and reduce variation), it should be clearly seen at least on one side of the slice section. If the cut was too rostral, make another section and check for the SCN under magnification.</li>
</ol>
4. Organotypic SCN Culture
<ol>
<li>In the petri dish lid filled with HBSS, dissect out the bilateral SCN as a square tissue (~ 1.5 mm each side) with a pair of sterile scalpels under a dissecting microscope. A small piece of the optic chiasm will remain attached to the explant, but no other nuclei should be included. Cut as close as possible without removing SCN tissue.</li>
<li>If appropriate for the planned experiment, the bilateral SCN can then be cut in half to obtain two unilateral SCN. One unilateral SCN can then be used as control (Fig 2a).</li>
<li>Fill 1200 &mu;L of the luciferin-medium into a &#8776; 35 mm Petri dish and place a culture membrane (Milli-CM 0.4 &mu;m, Millipore, Bedford, MA) on top of the liquid surface (2b). The culture medium volume is crucial, as the culture membrane should sit securely on the base of the culture dish, not float or rock in the medium 11. Make sure there are no air bubbles under the membrane. Avoid unnecessary light exposure when working with luciferin.</li>
<li>The explants are small and difficult to pick up. Therefore use a 1000 &mu;L pipette + tip to suck the SCN explant into the tip and press it out on the membrane. In case the explant is too big to easily fit into a 1000 &mu;L pipette tip (e.g. rat SCN; and mouse bilateral SCN) the tip may be cut with a sterile tool to create a wider opening. Discard excessive HBSS on the membrane with the pipette. For luciferase recording, place only one SCN/dish (Fig 2b) as the recording tubes detect all photons emitted from the dish and do not distinguish between signals from different tissues. </li>
<li>Seal the dish with a cover glass (&#8776; 40 mm, Menzel-Gl&auml;ser, Germany) and vacuum grease (Dow Corning Corp, USA)11. Make sure the seal is tight (Fig 2c). If not, seal with more grease.</li>
<li>Transfer the dishes to a 36-37&deg;C light-tight chamber and start PMT-recordings immediately.</li>
</ol>
5. Measurement of Luciferase Activity by Recording Bioluminescence
Luciferase-induced bioluminescence signals from the small SCN tissue are detected and amplified with photonmultiplier-tube (PMT) detector assemblies mounted inside a light tight chamber. The PMTs are normally positioned ~ 1-2 cm above the culture dishes 8. PMT recording setups can be custom made or are commercially available 11.
<ol>
<li>Place the dish under a PMT (the heat produced from the PMT will remove condensation on the cover glass) and close the chamber. Make sure the chamber is 100% light-tight as the dark count of PMTs used for SCN tissues is very low (less than 20 photons/min). The PMTs detect even the weakest light leakage.</li>
<li>Start the data acquisition, which is performed with software (for instance LumiCycle; Actimetrics Inc., Wilmette, IL, USA). Photon counts are integrated over 1-10 min intervals to get high resolution of the gene/protein expression.</li>
<li>After the recording is finished, the obtained circadian expression of the gene/protein can be analyzed with appropriate software (Origin, OriginLab, Northampton, MA, USA; ClockLab, Actimetrics; LumiCycle, Actimetrics Inc; Chrono, Till Roenneberg, University of Munich, Munich, Germany) to determine phase, period (time for one cycle) and amplitude of the rhythm. The peak expression of gene or protein is mostly used as reference point and is defined as the highest photon count during one cycle. The data can be smoothed before analyses of phase, period and amplitude, especially if the signal/noise ratio is low. The baseline sometimes changes and need to be subtracted before the analyses are performed.</li>
</ol>
6. Representative Results:
We here present the oscillatory PER2::LUC expression as a read out for viability and condition of the cultured tissue. Under optimal conditions, and if the tissue is alive, the PER2::LUC expression oscillates with a circadian rhythm as shown in figure 3. PER2 in the SCN is maximally expressed typically around Zeitgeber Time 12-13 (where ZT 12 represents lights off in a 12:12 hr light:dark cycle). The larger in size the live tissue is, the higher the photon count becomes. However, the size and thickness of organotypically cultured tissue should be kept small in order to keep the tissue viable, preferably not larger than 15 mm2 11 and not thicker than 500 &mu;m 7. The SCN, if dissected as described here, typically shows photon counts between 10.000-40.000/min if the tissue is sampled from a homozygous PER2::LUC animal. The amplitude of the oscillation in organotypic SCN cultures is typically very high during the first cycle as compared with the following cycles. It is not entirely clear why the first cycle has very high amplitude. One possible explanation is that a substantial portion of cells in the tissue may die shortly after initial cutting and culturing, thus not utilizing luciferin after the first cycle. The cutting procedure may also cause excessive excitation, which could amplify the luciferase signal during the first cycle.
Figure 3 shows luminescence traces from SCN slices and contains one trace (red) obtained from a slice that was not initially healthy. Dead or unhealthy tissues have low photon count baselines. (In addition, dead tissues often dissociate in the dish and cannot be removed from the membrane in one piece.)
The technique described in this report can beneficially be used in pharmacological experiments. Figure 4 shows a trace from a culture that we treated between day 1 and 2 with a HCN channel blocker (ZD7288, 10 &mu;M). As can be seen in the figure and as published previously 15, the blocker significantly reduced the amplitude of the circadian oscillation of PER2; however, after washing out with normal culture medium the oscillation came back, demonstrating that the blocker affected the molecular clock but the tissue was viable and healthy.

 <img src="/files/ftp_upload/2439/2439fig1.jpg" alt="Figure 1" /> 
Figure 1. Slicing procedure 
A) The head of a euthanized decapitated mouse with eyes and skin removed. B) The skull is removed with a micro rongeur tool. When using the tool, one must work upwards and never press down the brain with the tool. C) The brain shown upside down (ventral side up) without olfactory bulbs. The white optic chiasm with the two intact optic nerves can be seen. The suprachiasmatic nucleus (SCN, the borders marked with red) is located close to the optic chiasm. D) A coronally cut brain attached to the platform in the vibroslicer, at the level of SCN. E) Coronal brain section (250 &mu;m thick) containing the optic chiasm (OC), third ventricle (3V) and the bilateral suprachiasmatic nuclei (SCN).
 <img src="/files/ftp_upload/2439/2439fig2.jpg" alt="Figure 2" /> 
Figure 2. Organotypic tissue culturing. 
A) The two unilateral SCN nuclei (insert) dissected from the slice shown. B) Culture dish (35 mm Petri dish) with culture membrane, medium and explant, but without vacuum grease and cover glass. The medium (1.2 mL) can be seen as liquid between the dish and the membrane. One unilateral SCN nucleus is placed on the culture membrane. The whitest part of the small tissue is a piece of the optic chiasm (insert). C) The culture dish with its membrane and its SCN tissue, sealed with vacuum grease and a round cover glass.
 <img src="/files/ftp_upload/2439/2439fig3.jpg" alt="Figure 3" /> 
Figure 3. Bioluminescence recordings from healthy and non-healthy SCN tissue cultures. 
Examples of bioluminescence recordings of PERIOD2::LUCIFERASE (PER2::LUC) expression in suprachiasmatic nucleus (SCN) slices obtained from mice held in a 12h:12h light:dark cycle. The PER2::LUC protein oscillates with a circadian (~24 hr) variation in which the maximum expression of the protein occurs at Zeitgeber time 12-13. Thus, the phase of the gene rhythm is dependent on the light dark schedule in which the animal was kept before sacrificed. Typically, bioluminescence from unilateral SCN tissues dissected as described in the protocol shows photon counts between ~10.000-40.000/minute. The figure shows traces from one healthy (black) SCN culture, one non-healthy SCN culture (red) and one SCN culture that dried out (blue) after opening the sealed culture dish at day 4 and not re-sealing the dish properly (indicated by arrow).
 <img src="/files/ftp_upload/2439/2439fig4.jpg" alt="Figure 4" /> 
Figure 4. Bioluminescence recording during and after drug exposure. 
PER2::LUC expression in a culture before, during and after action of a HCN channel blocker (ZD7288, 10 &mu;M). The first arrow indicates the time when the blocker was added. The second arrow indicates washout, which was made by replacing the drug containing medium with conditioned control medium. Note the reduced amplitude after 2 days of drug exposure, lack of oscillation at day 4 and the quick recovery of the protein rhythm after washout.

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G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+brain+slice+chamber+designs+for+investigations+of+oxygen+deprivation+in+vitro.">Comparison of brain slice chamber designs for investigations of oxygen deprivation in vitro.</a> J Neurosci Methods. 81, 103-111 (1998).</li><li>Burgoon, P. W., Boulant, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temperature-sensitive+properties+of+rat+suprachiasmatic+nucleus+neurons.">Temperature-sensitive properties of rat suprachiasmatic nucleus neurons.</a> Am J Physiol Regul Integr Comp Physiol. 281, 706-715 (2001).</li><li>Nishide, S. Y., Honma, S., Honma, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+circadian+pacemaker+in+the+cultured+suprachiasmatic+nucleus+from+pup+mice+is+highly+sensitive+to+external+perturbation.">The circadian pacemaker in the cultured suprachiasmatic nucleus from pup mice is highly sensitive to external perturbation.</a> Eur J Neurosci. 27, 2686-2690 (2008).</li><li>Yoshikawa, T., Sellix, M., Pezuk, P., Menaker, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Timing+of+the+ovarian+circadian+clock+is+regulated+by+gonadotropins.">Timing of the ovarian circadian clock is regulated by gonadotropins.</a> Endocrinology. 150, 4338-4347 (2009).</li><li>Kohn, R. A., Dunlap, T. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calculation+of+the+buffering+capacity+of+bicarbonate+in+the+rumen+and+in+vitro.">Calculation of the buffering capacity of bicarbonate in the rumen and in vitro.</a> J Anim Sci. 76, 1702-1709 (1998).</li><li>Wilsbacher, L. D., Yamazaki, S., Herzog, E. D., Song, E. J., Radcliffe, L. A., Abe, M., Block, G., Spitznagel, E., Menaker, M., Takahashi, 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=Photic+and+circadian+expression+of+luciferase+in+mPeriod1-luc+transgenic+mice+in+vivo.">Photic and circadian expression of luciferase in mPeriod1-luc transgenic mice in vivo.</a> Proc Natl Acad Sci U S A. 99, 489-494 (2002).</li><li>Sellix, M. T., Currie, J., Menaker, M., Wijnen, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescence/luminescence+circadian+imaging+of+complex+tissues+at+single-cell+resolution.">Fluorescence/luminescence circadian imaging of complex tissues at single-cell resolution.</a> J Biol Rhythms. 25, 228-232 (2010).</li></ol>

No conflicts of interest declared.

Benefits and disadvantages with luciferase reporter technology
In contrast to ex vivo methods, such as RT-PCR, in situ hybridization and Western blot that require tissue sampling at many different time points (giving a low time resolution of normally 2-4 hrs depending on sampling frequency) in order to study diurnal variations in gene and protein expressions, the luciferase reporter technology allows high resolution (1-10 min) studies of circadian oscillations for many days in the same preparation. Thus, the number of used animals is minimized and detailed studies of effects on phase and period of the rhythm are feasible, which normally is not possible using conventional sampling techniques with low time resolution. However, although relative amplitude measurements of the rhythm are possible, it should be emphasized that reporter technology is not quantitative and can therefore not be used to measure amounts of transcribed genes or translated proteins.
The tissue recordings can be started and analyzed immediately after culturing, a great advantage for directly studying molecular effects of in vivo stress, in vivo light-induced phase shifts etc. The organotypic SCN culture allows long-term (weeks) pharmacological manipulation of adult brain tissue containing an intact mature synaptic network and the luciferase reporter technology allows stable recordings for many weeks. Thus, the tissue does not need to be neonatal or early postnatal in order to obtain healthy cultures, and in contrast to the acute slice chronic pharmacological treatments can be performed. In contrast to plasmid-reporter transfections in cell cultures, which do no lead to incorporation of DNA into the genome, the transgenic luciferase-animal models (as well as lenti-virus transfections) are favorable if epigenetic alterations are to be studied. It should in this context also be mentioned that luminescence imaging is nowadays possible with highly sensitive CCD cameras 11, allowing recordings even in single SCN neurons (~ 6-10 &mu;m) and other cell types, utilized by major laboratories studying circadian rhythms. Finally, several circadian investigators commonly use monitoring of molecular expression using other reporters, such as the Green Fluorescent Protein, in order to study clock gene/protein oscillations 16-19.
Aspects of slice thickness
The here recommended thicknesses of the SCN slice (200-300 &mu;m) could be reduced or increased if desired. However, although the tissue flattens out on the membrane after a few days in culture, it is not recommended to exceed 500 &mu;m thickness of the initially cut slice in order to preserve viability of the tissue 7. A slice thinner than 100 &mu;m is of mechanical and practical reasons difficult to handle. Because the SCN tissue is heterogeneous the thickness of the slice may affect the phase of the luciferase output signal, since different regions within the SCN (for instance the "core" versus the "shell", and dorsal versus ventral regions) oscillate with different phases 20 and differentially re-synchronize after phase shifts 21. The thickness of the slice also affects the baseline of photon count since more tissue emits a larger number of photons. The amplitude of molecular oscillations may in this way indirectly be affected by the size of the explant. For these reasons, control and treated cultures should always be of the same size, thickness and containing the same region of SCN in order to oscillate in the same phase and with the same amplitude. The two unilateral nuclei, on the other hand, oscillate in phase with each other and with similar amplitudes as long as the vibratome cut is horizontal and not angled (which in turn is dependent on that the separation cut between cerebellum and the two hemispheres is perpendicular to the table surface). An investigator who performs SCN culturing might experience that SCN cultures do not oscillate in phase with each other. Practicing and standardization of the slicing technique, i.e. the slices are always cut at the same rostro-caudal level of the SCN, will improve the outcome.
Critical steps
<ol>
<li>Oxygen supply is critical. 
Because the brain requires a lot of oxygen 22, the slicing procedure needs to be quick (The closer 3.1-3.10 is to ~5-6 minutes the better it is in order to keep the tissue healthy).</li>
<li>The SCN is sensitive to temperature changes and medium exchange. 
The SCN is temperature sensitive and big temperature changes can phase shift the pacemaker and cause experimental artifacts 23. If medium or solutions are exchanged or added during an ongoing experiment, for example when applying a drug after a couple of days in culture, the medium or solution needs to be pre-warmed to 36-37&deg;C (the same temperature as in the chamber) before addition. In pharmacological experiments that involve medium exchanges, it is important to always work with pre-conditioned cell culture medium. Addition/wash-out with fresh, unconditioned medium can potentially phase shift the molecular SCN rhythm 24 as well as other cell type cultures 25.</li>
<li>Medium composition. 
It is important to culture the tissue in correct pH and osmolality. The air-buffering medium contains a large amount of HEPES, which has an optimal buffer capacity in the range of pH 6.8-8.2 and is also appropriate for buffering the pH changes that may occur as a result of cell respiration. HEPES, on the other hand, is more sensitive to temperature changes as compared with sodium bicarbonate, which is included in small amounts in the culture medium. Sodium bicarbonate has a greater buffering capacity in the lower pH (5.1-7.1) range 26, thus appropriate in a CO2 atmosphere. However, increased amounts of sodium bicarbonate in the air-buffering medium (DMEM 2902) increases the osmolality significantly and can not be added without simultaneously diluting the medium, which in turn results in the incorrect composition of amino acids, vitamins and ions. 
Advice for preparing the medium:
<ol>
<li>Be precise and careful when mixing the medium.</li>
<li>Use only fresh or freshly thawed stock solutions.</li>
<li>In case of very high osmolality it is not worth diluting. More than 5-6%-diluted medium will most likely not work. Use D-glucose to increase osmolality if it is too low.</li>
</ol>
</li>
<li>Phase shifts due to preparation time 
The time of slice preparation is critical. Zero or minimal effect on phase is obtained if the slice procedure is performed between ZT 6-12 12. All slice dissections should be performed at the same phase of the diurnal or circadian cycle in order to minimize error due to phase variation between the preparations.</li>
</ol>
Possible modifications
<ol>
<li>For acute SCN electrophysiology, the slicing has to be performed with a Ringer (Artificial Cerebro Spinal Fluid; ACSF) buffer, saturated with oxygen (95% O2, 5% CO2) before the slicing starts. Further, oxygen needs to be added continuously to the buffer during the entire slicing procedure, as the electrical and synaptic activity is directly dependent on oxygen supply 22.</li>
<li>Horizontal and sagittal slices can also be successfully prepared and cultured of the SCN. The authors have experience with luciferase recordings in horizontal slices, which according to our experience give circadian oscillations with lower amplitude as compared with the coronal cut.</li>
<li>The SCN is approximately 1 mm long rostro-caudally and the slicing technique described here is targeted towards acquiring an SCN section in the central region. However, more than one SCN section can be obtained from mouse and rat brains, allowing the investigation of circadian gene/protein expression at more rostral versus caudal levels of the SCN nucleus.</li>
<li>Slice cultures from younger animals, postnatal pups or even embryos, can also be prepared. Please note that the brain and the skull in very young pups are very soft and sensitive. In addition, the optic nerve is thin and not fully developed. Take caution when dissecting the brain of pups so that the SCN region is not damaged. For instance, micro rongeur tools cannot be used to open the skull of mouse pups because they are too big. Fine scissors are enough to cut the soft tissues in young pups.</li>
<li>The cover glass based culture technique described here can also be used to image luminescence using a CCD/EM-CCD camera, also in single SCN neurons.</li>
<li>Other central nervous system regions and peripheral organ tissues from transgenic PER2::LUC animals can easily be obtained, cultured and analyzed in terms of molecular oscillations, as the PER2::LUC expression continues to oscillate for many days also in a number of other tissues and cell types 10. Most of these peripheral tissues, for instance liver, do not require a membrane for survival but are instead cultured directly bathed in the medium 11. Please note that phase shifting due to sectioning, culturing and medium exchange may not follow the same principles as for SCN.</li>
</ol>
Significance
In transgenic mouse and rat strains (mPER2::LUC; mPer1-luc 8, 10, 27) luciferase enzyme activity reflects protein or gene expression rhythms and can be assessed by bioluminescence recording. The luciferase-generated bioluminescence gives a weak signal but the background luminescence is close to zero, making this method advantageous. Moreover, because the luciferase molecule is unstable and rapidly degraded there is no phototoxicity, which can appear during long-term excitatory illumination 28. Taken together, these properties allow long-lasting experiments making the luciferase reporter technology highly advantageous in circadian research.

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		<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>B27 supplement 50x</td>
<td>&nbsp;</td>
<td>Invitrogen/Gibco</td>
<td>17504-044</td>
<td>&nbsp;</td>
<tr>
<td>Cover glasses</td>
<td>&nbsp;</td>
<td>Menzel-Gl&auml;ser</td>
<td>40#1</td>
<td>&#8776; 40 mm</td>
</tr>
<tr>
<td>Culture membranes</td>
<td>&nbsp;</td>
<td>Millipore</td>
<td>PICMORG50</td>
<td>0.4 &mu;m</td>
</tr>
<tr>
<td>DMEM low glucose w/o phenol red</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>D2902</td>
<td>Powder for 1L</td>
</tr>
<tr>
<td>Filter paper</td>
<td>&nbsp;</td>
<td>Whatman</td>
<td>1003 055</td>
<td>&#8776; 55 mm</td>
</tr>
<tr>
<td>Filtration set</td>
<td>&nbsp;</td>
<td>Corning</td>
<td>431097</td>
<td>Pore size 0.22 &mu;m
Polyethersulfone</td>
</tr>
<tr>
<td>Forceps </td>
<td>&nbsp;</td>
<td>Allgaier Instrumente</td>
<td>0203-7-PS</td>
<td>Dumant #7</td>
</tr>
<tr>
<td>HEPES</td>
<td>&nbsp;</td>
<td>Invitrogen/Gibco</td>
<td>15630-056</td>
<td>100 ml</td>
</tr>
<tr>
<td>HBSS 10x</td>
<td>&nbsp;</td>
<td>Invitrogen/Gibco </td>
<td>14065-049</td>
<td>500 ml</td>
</tr>
<tr>
<td>NaOCH3 7.5%</td>
<td>&nbsp;</td>
<td>Invitrogen/Gibco</td>
<td>25080-060</td>
<td>50 ml</td>
</tr>
<tr>
<td>Luciferin </td>
<td>&nbsp;</td>
<td>Promega</td>
<td>E1602</td>
<td>Beetle luciferin, potassium salt</td>
</tr>
<tr>
<td>Micro rongeur tool</td>
<td>&nbsp;</td>
<td>Allgaier Instrumente</td>
<td>332-097-140</td>
<td>&nbsp;</td>
<tr>
<td>PenStrep 10,000 U/ml</td>
<td>&nbsp;</td>
<td>Invitrogen/Gibco</td>
<td>15140-122</td>
<td>100 ml</td>
</tr>
<tr>
<td>Petri dishes</td>
<td>&nbsp;</td>
<td>Corning</td>
<td>430165</td>
<td>&#8776; 35 mm</td>
</tr>
<tr>
<td>PMT</td>
<td>&nbsp;</td>
<td>Hamamatsu</td>
<td>H9319-11MOD</td>
<td>&nbsp;</td>
<tr>
<td>Disposable scalpels</td>
<td>&nbsp;</td>
<td>Paragon</td>
<td>P503</td>
<td>Size 11</td>
</tr>
<tr>
<td>Vacuum filter</td>
<td>&nbsp;</td>
<td>Fisher </td>
<td>09-761-5</td>
<td>&nbsp;</td>
<tr>
<td>Vacuum grease</td>
<td>&nbsp;</td>
<td>Dow Corning</td>
<td>&nbsp;</td>
<td>50 g</td>
</tr>
<tr>
<td>Vibratome</td>
<td>&nbsp;</td>
<td>Campden Instruments</td>
<td>&nbsp;</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

slice preparation, organotypic tissue culturing and luciferase recording of clock gene activity in the suprachiasmatic nucleus

A central circadian (~24 hr) clock coordinating daily rhythms in physiology and behavior resides in the suprachiasmatic nucleus (SCN) located in the anterior hypothalamus. The clock is directly synchronized by light via the retina and optic nerve. Circadian oscillations are generated by interacting negative feedback loops of a number of so called "clock genes" and their protein products, including the Period (Per) genes. The core clock is also dependent on membrane depolarization, calcium and cAMP 1. The SCN shows daily oscillations in clock gene expression, metabolic activity and spontaneous electrical activity. Remarkably, this endogenous cyclic activity persists in adult tissue slices of the SCN 2-4. In this way, the biological clock can easily be studied in vitro, allowing molecular, electrophysiological and metabolic investigations of the pacemaker function.
The SCN is a small, well-defined bilateral structure located right above the optic chiasm 5. In the rat it contains ~8.000 neurons in each nucleus and has dimensions of approximately 947 &mu;m (length, rostrocaudal axis) x 424 &mu;m (width) x 390 &mu;m (height) 6. To dissect out the SCN it is necessary to cut a brain slice at the specific level of the brain where the SCN can be identified. Here, we describe the dissecting and slicing procedure of the SCN, which is similar for mouse and rat brains. Further, we show how to culture the dissected tissue organotypically on a membrane 7, a technique developed for SCN tissue culture by Yamazaki et al. 8. Finally, we demonstrate how transgenic tissue can be used for measuring expression of clock genes/proteins using dynamic luciferase reporter technology, a method that originally was used for circadian measurements by Geusz et al. 9. We here use SCN tissues from the transgenic knock-in PERIOD2::LUCIFERASE mice produced by Yoo et al. 10. The mice contain a fusion protein of PERIOD (PER) 2 and the firefly enzyme LUCIFERASE. When PER2 is translated in the presence of the substrate for luciferase, i.e. luciferin, the PER2 expression can be monitored as bioluminescence when luciferase catalyzes the oxidation of luciferin. The number of emitted photons positively correlates to the amount of produced PER2 protein, and the bioluminescence rhythms match the PER2 protein rhythm in vivo 10. In this way the cyclic variation in PER2 expression can be continuously monitored real time during many days. The protocol we follow for tissue culturing and real-time bioluminescence recording has been thoroughly described by Yamazaki and Takahashi 11.

Watch this Scientific Journal Video about Slice Preparation, Organotypic Tissue Culturing and Luciferase Recording of Clock Gene Activity in the Suprachiasmatic Nucleus at JoVE.com

Slice Preparation, Organotypic Tissue Culturing and Luciferase Recording of Clock Gene Activity in the Suprachiasmatic Nucleus

The procedure of preparing slices containing the adult mouse hypothalamic suprachiasmatic nucleus (SCN), and a rapid way to culture the SCN tissue in organotypic culture condition, are reported. Further, the measurement of oscillatory clock gene protein expression using dynamic luciferase reporter technology is described.

A central circadian (~24 hr) clock coordinating daily rhythms in physiology and behavior resides in the suprachiasmatic nucleus (SCN) located in the ...

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Research

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Neuroscience

The Organotypic Hippocampal Slice Culture Model for Examining Neuronal Injury

Organotypic hippocampal slice culture is an in vitro method to examine mechanisms of neuronal injury in which the basic architecture and composition of the hippocampus is relatively preserved 1. The organotypic culture system allows for the examination of neuronal, astrocytic and microglial effects, but as an ex vivo preparation, does not address effects of blood flow, or recruitment of peripheral inflammatory cells. To that end, this culture method is frequently used to examine excitotoxic and hypoxic injury to pyramidal neurons of the hippocampus, but has also been used to examine the inflammatory response. Herein we describe the methods for generating hippocampal slice cultures from postnatal rodent brain, administering toxic stimuli to induce neuronal injury, and assaying and quantifying hippocampal neuronal death.

The organoptypic hippocampal slice culture model is an in vitro model used to examine neuronal injury in a variety of paradigms. In this article, we describe the methods for generating slice cultures and quantifying neuronal injury.

Organotypic hippocampal slice culture is an in vitro method to examine mechanisms of neuronal injury in which the basic architecture and composition of ...

Organotypic Hippocampal Slice Cultures

The hippocampus, a component of the limbic system, plays important roles in long-term memory and spatial navigation 1. Hippocampal neurons can modify the strength of their connections after brief periods of strong activation. This phenomenon, known as long-term potentiation (LTP) can last for hours or days and has become the best candidate mechanism for learning and memory 2. In addition, the well defined anatomy and connectivity of the hippocampus 3 has made it a classical model system to study synaptic transmission and synaptic plasticity4.
As our understanding of the physiology of hippocampal synapses grew and molecular players became identified, a need to manipulate synaptic proteins became imperative. Organotypic hippocampal cultures offer the possibility for easy gene manipulation and precise pharmacological intervention but maintain synaptic organization that is critical to understanding synapse function in a more naturalistic context than routine culture dissociated neurons methods.
Here we present a method to prepare and culture hippocampal slices that can be easily adapted to other brain regions. This method allows easy access to the slices for genetic manipulation using different approaches like viral infection 5,6 or biolistics 7. In addition, slices can be easily recovered for biochemical assays 8, or transferred to microscopes for imaging 9 or electrophysiological experiments 10.

We describe a method to prepare organotypic hippocampal slices that can be easily adapted to other brain regions. Brain slices are laid on porous membranes and culture media is allowed to form an interface. This method preserves the gross architecture of the hippocampus for up to 2 weeks in culture.

The hippocampus, a component of the limbic system, plays important roles in long-term memory and spatial navigation 1. Hippocampal neurons can modify the ...

Whole Cell Recording from an Organotypic Slice Preparation of Neocortex

We have been studying the expression and functional roles of voltage-gated potassium channels in pyramidal neurons from rat neocortex. Because of the lack of specific pharmacological agents for these channels, we have taken a genetic approach to manipulating channel expression. We use an organotypic culture preparation (16) in order to maintain cell morphology and the laminar pattern of cortex. We typically isolate acute neocortical slices at postnatal days 8-10 and maintain the slices in culture for 3-7 days. This allows us to study neurons at a similar age to those in our work with acute slices and minimizes the development of exuberant excitatory connections in the slice. We record from visually-identified pyramidal neurons in layers II/III or V using infrared illumination (IR-) and differential interference contrast microscopy (DIC) with whole cell patch clamp in current- or voltage-clamp. We use biolistic (Gene gun) transfection of wild type or mutant potassium channel DNA to manipulate expression of the channels to study their function. The transfected cells are easily identified by epifluorescence microscopy after co-transfection with cDNA for green fluorescent protein (GFP). We compare recordings of transfected cells to adjacent, untransfected neurons in the same layer from the same slice.

This is a protocol to prepare and maintain a neocortical slice preparation in organotypic culture for the purpose of making electrical recordings from pyramidal neurons.

We have been studying the expression and functional roles of voltage-gated potassium channels in pyramidal neurons from rat neocortex.  Because of the ...

Imaging Pheromone Sensing in a Mouse Vomeronasal Acute Tissue Slice Preparation

Peter Karlson and Martin L&uuml;scher used the term pheromone for the first time in 19591 to describe chemicals used for intra-species communication. Pheromones are volatile or non-volatile short-lived molecules2 secreted and/or contained in biological fluids3,4, such as urine, a liquid known to be a main source of pheromones3. Pheromonal communication is implicated in a variety of key animal modalities such as kin interactions5,6, hierarchical organisations3 and sexual interactions7,8 and are consequently directly correlated with the survival of a given species9,10,11. In mice, the ability to detect pheromones is principally mediated by the vomeronasal organ (VNO)10,12, a paired structure located at the base of the nasal cavity, and enclosed in a cartilaginous capsule. Each VNO has a tubular shape with a lumen13,14 allowing the contact with the external chemical world. The sensory neuroepithelium is principally composed of vomeronasal bipolar sensory neurons (VSNs)15. Each VSN extends a single dendrite to the lumen ending in a large dendritic knob bearing up to 100 microvilli implicated in chemical detection16. Numerous subpopulations of VSNs are present. They are differentiated by the chemoreceptor they express and thus possibly by the ligand(s) they recognize17,18. Two main vomeronasal receptor families, V1Rs and V2Rs19,20,21,22, are composed respectively by 24023 and 12024 members and are expressed in separate layers of the neuroepithelium. Olfactory receptors (ORs)25 and formyl peptide receptors (FPRs)26,27 are also expressed in VSNs.
Whether or not these neuronal subpopulations use the same downstream signalling pathway for sensing pheromones is unknown. Despite a major role played by a calcium-permeable channel (TRPC2) present in the microvilli of mature neurons28 TRPC2 independent transduction channels have been suggested6,29. Due to the high number of neuronal subpopulations and the peculiar morphology of the organ, pharmacological and physiological investigations of the signalling elements present in the VNO are complex.
Here, we present an acute tissue slice preparation of the mouse VNO for performing calcium imaging investigations. This physiological approach allows observations, in the natural environment of a living tissue, of general or individual subpopulations of VSNs previously loaded with Fura-2AM, a calcium dye. This method is also convenient for studying any GFP-tagged pheromone receptor and is adaptable for the use of other fluorescent calcium probes. As an example, we use here a VG mouse line30, in which the translation of the pheromone V1rb2 receptor is linked to the expression of GFP by a polycistronic strategy.

In mice, the ability to detect pheromones is principally mediated by the vomeronasal organ (VNO). Here, an acute tissue slice preparation of VNO for performing calcium imaging is described. This physiological approach allows observations of subpopulations and/or individual neurons in a living tissue and is convenient for receptor-ligand identification.

Peter Karlson and Martin L&uuml;scher used the term pheromone for the first time in 19591 to describe chemicals used for intra-species communication. ...

The Analysis of Purkinje Cell Dendritic Morphology in Organotypic Slice Cultures

Purkinje cells are an attractive model system for studying dendritic development, because they have an impressive dendritic tree which is strictly oriented in the sagittal plane and develops mostly in the postnatal period in small rodents 3. Furthermore, several antibodies are available which selectively and intensively label Purkinje cells including all processes, with anti-Calbindin D28K being the most widely used. For viewing of dendrites in living cells, mice expressing EGFP selectively in Purkinje cells 11 are available through Jackson labs. Organotypic cerebellar slice cultures cells allow easy experimental manipulation of Purkinje cell dendritic development because most of the dendritic expansion of the Purkinje cell dendritic tree is actually taking place during the culture period 4. We present here a short, reliable and easy protocol for viewing and analyzing the dendritic morphology of Purkinje cells grown in organotypic cerebellar slice cultures. For many purposes, a quantitative evaluation of the Purkinje cell dendritic tree is desirable. We focus here on two parameters, dendritic tree size and branch point numbers, which can be rapidly and easily determined from anti-calbindin stained cerebellar slice cultures. These two parameters yield a reliable and sensitive measure of changes of the Purkinje cell dendritic tree. Using the example of treatments with the protein kinase C (PKC) activator PMA and the metabotropic glutamate receptor 1 (mGluR1) we demonstrate how differences in the dendritic development are visualized and quantitatively assessed. The combination of the presence of an extensive dendritic tree, selective and intense immunostaining methods, organotypic slice cultures which cover the period of dendritic growth and a mouse model with Purkinje cell specific EGFP expression make Purkinje cells a powerful model system for revealing the mechanisms of dendritic development.

We present a protocol that permits to view and to quantitatively asses the morphology of the dendritic tree of individual Purkinje cells grown in organotypic cerebellar slice cultures. This protocol is intended to promote studies on the mechanisms of Purkinje cell dendritic development.

Purkinje cells are an attractive model system for studying dendritic development, because they have an impressive dendritic tree which is strictly ...

Recording and Analysis of Circadian Rhythms in Running-wheel Activity in Rodents

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, including rats, hamsters, and mice, are active during the night and relatively inactive during the day. Many other behavioral and physiological measures also exhibit daily rhythms, but in rodents, running-wheel activity serves as a particularly reliable and convenient measure of the output of the master circadian clock, the suprachiasmatic nucleus (SCN) of the hypothalamus. In general, through a process called entrainment, the daily pattern of running-wheel activity will naturally align with the environmental light-dark cycle (LD cycle; e.g. 12 hr-light:12 hr-dark). However circadian rhythms are endogenously generated patterns in behavior that exhibit a ~24 hr period, and persist in constant darkness. Thus, in the absence of an LD cycle, the recording and analysis of running-wheel activity can be used to determine the subjective time-of-day. Because these rhythms are directed by the circadian clock the subjective time-of-day is referred to as the circadian time (CT). In contrast, when an LD cycle is present, the time-of-day that is determined by the environmental LD cycle is called the zeitgeber time (ZT).
Although circadian rhythms in running-wheel activity are typically linked to the SCN clock6-8, circadian oscillators in many other regions of the brain and body9-14 could also be involved in the regulation of daily activity rhythms. For instance, daily rhythms in food-anticipatory activity do not require the SCN15,16 and instead, are correlated with changes in the activity of extra-SCN oscillators17-20. Thus, running-wheel activity recordings can provide important behavioral information not only about the output of the master SCN clock, but also on the activity of extra-SCN oscillators. Below we describe the equipment and methods used to record, analyze and display circadian locomotor activity rhythms in laboratory rodents.

Circadian rhythms in voluntary wheel-running activity in mammals are tightly coupled to the molecular oscillations of a master clock in the brain. As such, these daily rhythms in behavior can be used to study the influence of genetic, pharmacological, and environmental factors on the functioning of this circadian clock.

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, ...

Simultaneous Electrophysiological Recording and Calcium Imaging of Suprachiasmatic Nucleus Neurons

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in regulating the intracellular calcium concentration. Changing environmental conditions, such as the light-dark cycle, can modify neuronal and neural network activity and the expression of a family of circadian clock genes within the suprachiasmatic nucleus (SCN), the location of the master circadian clock in the mammalian brain. Excitatory synaptic transmission leads to an increase in the postsynaptic Ca2+ concentration that is believed to activate the signaling pathways that shifts the rhythmic expression of circadian clock genes. Hypothalamic slices containing the SCN were patch clamped using microelectrodes filled with an internal solution containing the calcium indicator bis-fura-2. After a seal was formed between the microelectrode and the SCN neuronal membrane, the membrane was ruptured using gentle suction and the calcium probe diffused into the neuron filling both the soma and dendrites. Quantitative ratiometric measurements of the intracellular calcium concentration were recorded simultaneously with membrane potential or current. Using these methods it is possible to study the role of changes of the intracellular calcium concentration produced by synaptic activity and action potential firing of individual neurons. In this presentation we demonstrate the methods to simultaneously record electrophysiological activity along with intracellular calcium from individual SCN neurons maintained in brain slices.

Procedures are described to perform simultaneous recordings of membrane potential or current and changes of intracellular calcium concentration. Suprachiasmatic nucleus neurons are filled with the calcium indicator bis-fura-2 using a patch clamp electrode in the whole cell patch clamp configuration.

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in ...

Organotypic Slice Cultures to Study Oligodendrocyte Dynamics and Myelination

NG2 expressing cells (polydendrocytes, oligodendrocyte precursor cells) are the fourth major glial cell population in the central nervous system. During embryonic and postnatal development they actively proliferate and generate myelinating oligodendrocytes. These cells have commonly been studied in primary dissociated cultures, neuron cocultures, and in fixed tissue. Using newly available transgenic mouse lines slice culture systems can be used to investigate proliferation and differentiation of oligodendrocyte lineage cells in both gray and white matter regions of the forebrain and cerebellum. Slice cultures are prepared from early postnatal mice and are kept in culture for up to 1 month. These slices can be imaged multiple times over the culture period to investigate cellular behavior and interactions. This method allows visualization of NG2 cell division and the steps leading to oligodendrocyte differentiation while enabling detailed analysis of region-dependent NG2 cell and oligodendrocyte functional heterogeneity. This is a powerful technique that can be used to investigate the intrinsic and extrinsic signals influencing these cells over time in a cellular environment that closely resembles that found in vivo.

A technique to study NG2 cells and oligodendrocytes using a slice culture system of the forebrain and cerebellum is described. This method allows examination of the dynamics of proliferation and differentiation of cells within the oligodendrocyte lineage where the extracellular environment can be easily manipulated while maintaining tissue cytoarchitecture.

NG2 expressing cells (polydendrocytes, oligodendrocyte precursor cells) are the fourth major glial cell population in the central nervous system. During ...

The Analysis of Neurovascular Remodeling in Entorhino-hippocampal Organotypic Slice Cultures

Ischemic brain injury is among the most common and devastating conditions compromising proper brain function and often leads to persisting functional deficits in the affected patients. Despite intensive research efforts, there is still no effective treatment option available that reduces neuronal injury and protects neurons in the ischemic areas from delayed secondary death. Research in this area typically involves the use of elaborate and problematic animal models. Entorhino-hippocampal organotypic slice cultures challenged with oxygen and glucose deprivation (OGD) are established in&#160;vitro models which mimic cerebral ischemia. The novel aspect of this study is that changes of the brain blood vessels are studied in addition to neuronal changes and the reaction of both the neuronal compartment and the vascular compartment can be compared and correlated. The methods presented in this protocol substantially broaden the potential applications of the organotypic slice culture approach. The induction of OGD or hypoxia alone can be applied by rather simple means in organotypic slice cultures and leads to reliable and reproducible damage in the neural tissue. This is in stark contrast to the complicated and problematic animal experiments inducing stroke and ischemia in vivo. By broadening the analysis to include the study of the reaction of the vasculature could provide new ways on how to preserve and restore brain functions. The slice culture approach presented here might develop into an attractive and important tool for the study of ischemic brain injury and might be useful for testing potential therapeutic measures aimed at neuroprotection.

A protocol for entorhino-hippocampal organotypic slice cultures, which allows reproducing many aspects of ischemic brain injury, is presented. By studying changes of the neurovasculature in addition to changes in the neurons, this protocol is a versatile tool to study plastic changes in neural tissue after injury.

Ischemic brain injury is among the most common and devastating conditions compromising proper brain function and often leads to persisting functional ...

Direct-current Stimulation and Multi-electrode Array Recording of Seizure-like Activity in Mice Brain Slice Preparation

Cathodal transcranial direct-current stimulation (tDCS) induces suppressive effects on drug-resistant seizures. To perform effective actions, the stimulation parameters (e.g., orientation, field strength, and stimulation duration) need to be examined in mice brain slice preparations. Testing and arranging the orientation of the electrode relative to the position of the mice brain slice are feasible. The present method preserves the thalamocingulate pathway to evaluate the effect of DCS on anterior cingulate cortex seizure-like activities. The results of the multichannel array recordings indicated that cathodal DCS significantly decreased the amplitude of the stimulation-evoked responses and duration of 4-aminopyridine and bicuculline-induced seizure-like activity. This study also found that cathodal DCS applications at 15 min caused long-term depression in the thalamocingulate pathway. The present study investigates the effects of DCS on thalamocingulate synaptic plasticity and acute seizure-like activities. The current procedure can test the optimal stimulation parameters including orientation, field strength, and stimulation duration in an in vitro mouse model. Also, the method can evaluate the effects of DCS on cortical seizure-like activities at both the cellular and network levels.

Studies have shown that cathodal transcranial direct-current stimulation can produce suppressive effects on drug-resistant seizures. In this study, an in vitro experimental setup was devised in which the direct-current stimulation and multielectrode array recording of seizure-like activity were evaluated in mice brain slice preparation. The direct-current stimulation parameters were evaluated.