Work in our lab is supported by the Dr. Miriam and Sheldon G. Adelson Medical Research Foundation and the Israeli Association for Fighting the A-T Disease.

See Table of Materials for details of materials, equipment, and antibodies. Animal procedures were employed under the ethical guidelines of Tel Aviv University&#39;s Ethics Committee after approval. The procedure is carried out on 10-day old mouse pups, regardless of their sex. If needed, genotyping is performed on the previous day using a tail biopsy and standard methods. Solutions are sterile and stored at 4 &#176;C unless indicated otherwise; see Table 1.
1. Preparation of cultures
NOTE: Conduct the procedure within a sterile environment and disinfect the work surface in advance using 70% ethanol.
<ol>
	<li>Add 1 mL of BME to each well of 6-well plates. Using sterile forceps, place a single cell culture insert into each well and incubate the plates in a 37 &#176;C/5% CO2 incubator for a minimum of 2 h prior to the dissection procedure.</li>
	<li>Set up a dissection area within a biological hood, containing the tissue chopper, 30 mm culture plates, a binocular microscope, and a light source.</li>
	<li>Immerse the surgical tools in 70% ethanol throughout the process. Use a urinary cup filled with 70% ethanol for immersion, and a separate urinary cup with 1x PBS to wash off the ethanol before use.</li>
	<li>For each dissection (i.e., for each cerebellum), prepare two 30 mm culture plates filled with cold dissection medium and leave them on ice.</li>
	<li>Spray the animal with 70% ethanol.</li>
	<li>Using sharp scissors, swiftly decapitate the animal. To expose the brain, make a small incision with scissors away from the cerebellum. Using tweezers with rounded ends, gently peel off the skull. Remove the skull in small pieces to avoid damaging the cerebellar tissue.</li>
	<li>Immediately transfer the brain into cold dissection medium or directly remove the cerebellum from within the skull. To do this, expose the cerebellum and separate the cerebellar cortex from the brainstem by disconnecting the three pairs of peduncles, using a fine curved iris spatula. To identify the peduncles, view from rostral to caudal: the paired superior cerebellar peduncles connect the midbrain to each side of the cerebellum; the paired middle cerebellar peduncles connect the pons to each side of the cerebellum; and the paired inferior cerebellar peduncles connect to the medulla to each side of to the cerebellum. Once the cerebellum is visible, separate it from the cerebrum using the same spatula: slide the spatula between the cerebellum, the superior colliculus, and the brainstem. 
	NOTE: Do not make a special effort to remove the meninges as they readily detach after the cerebellum is sliced.</li>
	<li>Set the tissue chopper to a slice thickness of 350 &#181;m.</li>
	<li>Prior to positioning the cerebellum on the chopper&#39;s base, remove excess dissecting buffer from the tissue. This will improve the tissue&#39;s adhesion to the chopper&#39;s base.</li>
	<li>Position the cerebellum on the cutting platform in a vertical orientation and carry out parasagittal slicing at a slice thickness of 350 &#181;m. Gently separate the slices using a fine iris spatula and put them into cold dissection medium.</li>
	<li>Transfer the sliced tissue back to a fresh cold dissection buffer. Use a 1,000 &#181;L tip to delicately guide the tissue slices into the dish containing cold dissection buffer.</li>
	<li>Under the binocular use two curved iris spatulas to carefully separate the tissue slices from each other. For cultures, use slices derived from the cerebellar vermis situated in the medial cortico-nuclear zone of the organ.</li>
	<li>Transfer each tissue slice onto a culture insert using a wide spatula and move each insert carrying a tissue slice into a well in the 6-well plates containing the pre-warmed medium. Leave the plates in the incubator.</li>
	<li>Replace the medium every other day: aspirate the used medium using a sterile glass pipette, and with&#160;a 5 mL serological pipette, add 1 mL of fresh MBE with the pipette&#39;s tip leaning against the well&#39;s wall. Make sure not to direct the medium flow onto the tissue. 
	NOTE: The tissue slice will exhibit an inflammation-like appearance for the first 5-7 days in culture and will be ready for experiments as soon as this phenomenon disappears and up to 2 weeks after culture establishment.</li>
</ol>
2. Treatment with DNA damaging agents, fixation, staining, and microscopic imaging
<ol>
	<li>Add DNA-damaging chemicals directly to the culture medium at appropriate final concentrations, for predetermined durations. After the exposure period, remove the chemicals by replacing the medium with fresh culture medium.</li>
	<li>If the DDR readout involves PAR staining, add an inhibitor of poly(ADP-ribose) glycohydrolase (PARG) to the culture medium at a final concentration of 10 &#181;M, 30 min prior to fixation. 
	NOTE: This treatment is critical as the PAR modification is short-lived and continuously degraded by PARG.</li>
	<li>Immediately prior to fixation, aspirate the medium and replace it with 1 mL of 0.1 M phosphate buffer at room temperature. 
	NOTE: Cold buffer might cause detachment of the tissue slices from the insert.</li>
	<li>Immediately fix the slices by replacing the buffer with precooled (-20 &#176;C) acetone:methanol mix (1:1) such that the tissue slice is submerged in this mix&#160;and leave for 20 min at -20 &#176;C. 
	NOTE: It is possible to stop at this stage and proceed with immunostaining later. To do so, wash the tissues 3x with 0.1 M phosphate buffer precooled to 4 &#176;C, making sure the tissue is submerged in the buffer. Tissues can be stored now in 0.1M phosphate buffer for up to 2 weeks at 4 &#176;C.</li>
	<li>Transfer the tissue slice into a well in a 48-well plate.</li>
	<li>Use a scalpel and a cutting board to remove the margins of the insert, cutting around the tissue slice. Place each slice in a well of a 48-well plate. Add 100 &#181;L of 0.1 M phosphate buffer.</li>
	<li>For permeabilization, aspirate the buffer, add to the well 100 &#181;L of 0.1 M phosphate buffer containing 1% Triton X-100, and leave for 5 min at room temperature.</li>
	<li>Aspirate the solution and add 100 &#181;L of blocking solution (see Table 1) per well. Shake gently at room temperature for 2 h.</li>
	<li>During the blocking step, dilute the primary antibody or anti-PAR reagent (1:800) in the blocking solution.</li>
	<li>Remove the blocking solution (no need to wash) and add 100 &#181;L of primary antibody mix to each well. Apply gentle shaking for 2 h at room temperature.</li>
	<li>Dilute the secondary antibodies in 0.1 M phosphate buffer containing 0.2% Triton X-100 (1:500) while minimizing their exposure to light.</li>
	<li>Aspirate the primary antibody solution and wash the slices 3 x 5 min with 0.1 M phosphate buffer containing 0.2% Triton X-100, at room temperature, with gentle shaking.</li>
	<li>Replace the wash solution with 100 &#181;L of secondary antibody solution (1:500 dilution). Continue gentle shaking for 2 h at room temperature in the dark (cover the plates with aluminum foil).</li>
	<li>To prepare a DAPI solution, dilute the stock solution (2 mg/mL) to 1 &#181;g/mL in 1x PBS.</li>
	<li>Twenty minutes before the end of the 2 h incubation with the secondary antibody, add 20 &#181;L of the final DAPI solution to each well and continue shaking for 20 min. Aspirate the liquid and wash with the wash solution for 3 x 5 min with gentle shaking.</li>
	<li>For mounting, place the tissue on a microscope glass slide with the tissue facing up, add mounting medium, and cover with a cover glass. Place the slides on a flat surface and let them dry overnight in the dark, at room temperature (avoid using mounting medium containing DAPI to prevent blurriness).</li>
	<li>Store the slides at 4 &#176;C in a lightproof box.</li>
	<li>Capture microscopic images using a confocal microscope and appropriate filters.</li>
</ol>

Cerebellar Purkinje Cells: DNA Damage Response in Organotypic Cultures

The authors declare no conflicts of interest related to this study.

General comments 
The major advantage of the organotypic culture system is that it facilitates studies using the cerebellar cortex tissue, preserving its structural organization for several weeks in the culture dish setup. This system is useful for conducting in-depth morphological analyses of Purkinje cells (PCs), including detailed examinations of dendritic spines and ultrastructural features52,53,54...

The integrity of cellular DNA is constantly under threat from DNA damaging agents, predominantly metabolic by-products like reactive oxygen species, which inflict tens of thousands of DNA lesions per cell daily1. The persistent upkeep of genome stability is essential for cellular homeostasis2,3. The cornerstone of this maintenance is the DNA damage response (DDR) - an intricate, layered signaling network that initiates specific DNA repair pathways while carefully adjusting many other cellular processes4,5. Deficits in the DDR are commonly manifested as &#39;genome instability syndromes,&#39; marked by chromosomal instability, progressive tissue deterioration, impaired growth or development, a predisposition to cancer, and heightened sensitivity to particular DNA-damaging agents6,7,8,9. Notably, neurodegeneration, which often includes cerebellar atrophy, is a distinct feature of many genome instability syndromes7,10,11,12.
The autosomal recessive disorder, ataxia-telangiectasia (A-T), is a well-documented example of a genome instability disorder13,14,15. This condition arises from null mutations in the ATM (A-T, mutated) gene, responsible for coding the pivotal protein kinase, ATM, which is known primarily as a DDR mobilizer in response to DNA double-strand breaks (DSBs)16,17. A-T manifests as a multisystem disorder, predominantly characterized by progressive cerebellar degeneration, leading to acute motor impairments, immunodeficiency, gonadal atrophy, cancer predisposition, and extreme sensitivity to ionizing radiation. Cultured cells from individuals with A-T show chromosomal instability and increased sensitivity to genotoxic agents, especially those causing DSBs15,18,19. Importantly, ATM also plays a role in repairing other DNA lesions, underscoring its broad significance in maintaining genome stability20,21,22.
Despite thorough research into ATM&#39;s numerous roles, the specific mechanism leading to cerebellar degeneration in A-T remains a topic of active debate, with various models proposed to elucidate this process23,24,25,26,27,28,29,30. Our model28 suggests that cerebellar degeneration in A-T patients begins with the dysfunction and eventual loss of Purkinje cells (PCs). Considering ATM&#39;s critical role in preserving genome stability in the face of ongoing DNA damage, PCs are particularly vulnerable to the absence of ATM. We attribute this vulnerability to the combination of their high metabolic activity, distinctive chromatin structure, and extensive transcriptional activity. Ultimately, it is suggested that the loss of PC function, and hence, their degeneration, is due to the stochastic, functional inactivation of genes, a consequence of producing defective transcripts28.
The study of PC biology in the laboratory is impeded by challenges in cultivating isolated PCs, as these cells rely heavily on their natural milieu and neighboring cells for survival and function, rendering them incompatible with dissociated culture growth. Nonetheless, PCs can remain viable for extended periods in tissue slice cultures. Cerebellar organotypic cultures, which are tissue slices typically derived from rodent cerebella, maintain the tissue&#39;s structural organization and support various experimental manipulations analogous to those possible with cultured cells. Therefore, these cultures allow for cerebellar studies within a controlled setting31,32,33,34,35,36,37,38,39,40,41,42,43. Specifically, in the context of A-T, murine cerebellar organotypic cultures have proven to be instrumental in exploring the DDR in Atm-deficient PCs40,41,42,43. While Atm-deficient mice display only a subtle cerebellar phenotype44,45,46,47, presumably due to differences between human and mouse cerebellar physiology, the assumption is that ATM&#39;s roles are largely conserved across these species. This notion is supported by our observations that the deficient response to DNA DSBs in Atm-/- murine PCs aligns with that observed in other murine and human ATM/Atm-deficient cell types40,41,42,43.
A limitation in analyzing PC responses to various stimuli or stresses within organotypic cultures is the necessity to rely on microscopic imaging for readouts. The DDR is usually studied using bulk biochemical readouts, although common immunofluorescent markers are utilized as well, such as following the dynamics of formation and resolution of nuclear foci of phosphorylated histone H2AX (&#947;H2AX) and the 53BP1 protein, which are considered indicators of DSBs48,49. A broader measure is the fluorescent imaging of poly(ADP-ribose) (PAR) chain formation on proteins, a rapid and robust early DNA damage response, particularly to strand breaks7,50. We modified the protocol by Komulainen et al.51 for PAR staining in cerebellar organotypic cultures. We observed a pronounced PAR response in the sizeable nuclei of PCs. Presented here is our refined protocol for establishing murine cerebellar organotypic cultures and for visualizing the PAR response under genotoxic stress.

<table><tbody><tr><td>&lt;strong&gt;Reagent and Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>150 x 30 cm Petri dish</td><td>Corning Inc.</td><td>3261</td><td></td></tr><tr><td>35 x 10 mm Petri dish</td><td>Corning Inc.</td><td>3294</td><td></td></tr><tr><td>Acetone</td><td>Invitrogen</td><td>25030081</td><td></td></tr><tr><td>Acetone</td><td>Sigma-Aldrich</td><td>270725-1L</td><td></td></tr><tr><td>Adhesive microscope slides</td><td>Marienfeld</td><td>48187</td><td>76 x 26 x 1 mm</td></tr><tr><td>Blades for chopper</td><td>TED PELLA</td><td>Model TC752-1</td><td>&amp;nbsp;Style Razor Blades</td></tr><tr><td>BME</td><td>GIBCO</td><td>41010-026 500ml.&amp;nbsp;</td><td>No L-Glut</td></tr><tr><td>Cell culture inserts</td><td>Millipore</td><td>PICM0RG50</td><td></td></tr><tr><td>D-Glucose</td><td>Biological Indo.</td><td>02-015-1A</td><td></td></tr><tr><td>HBSS w/o phenol red</td><td>Sigma-Aldrich</td><td>H-1138-500L</td><td>No Ca+,No Mg+</td></tr><tr><td>HBSS with phenol red</td><td>Sartorius</td><td>02-015-1A</td><td></td></tr><tr><td>Horse serum&amp;nbsp;</td><td>GIBCO</td><td>26050088</td><td>heat inactivated</td></tr><tr><td>Iris forceps</td><td>CellPath</td><td>GZX-0100-00A</td><td>130mm blunt serrated</td></tr><tr><td>Iris spatula</td><td>F.S.T</td><td>10092-12</td><td></td></tr><tr><td>L-Glutamine (200 mM)</td><td>Gibco</td><td>41010026</td><td></td></tr><tr><td>Methanol</td><td>Sigma-Aldrich</td><td>322412-1L</td><td></td></tr><tr><td>Methanol&amp;nbsp;</td><td>Sigma-Aldrich</td><td>G5767</td><td></td></tr><tr><td>Microscope</td><td>Olympus</td><td></td><td></td></tr><tr><td>Mounting Medium without</td><td>GRIGENE. Ltd, Israel</td><td>K239320A</td><td></td></tr><tr><td>Paraquat</td><td>Sigma-Aldrich</td><td>856177-250MG</td><td>Paraquat dichloride (Methyl viologen)</td></tr><tr><td>PARG- Poly(ADP-Ribose) Glycohydrolase inhibitor</td><td>Tocris Bioscience, United Kingdom</td><td>PDD 00017273</td><td>light sensitive&amp;nbsp;</td></tr><tr><td>Phosphate buffer</td><td>Biological Indo.</td><td>02-018-1A</td><td></td></tr><tr><td>Scissors, skin-tissue</td><td></td><td></td><td></td></tr><tr><td>Six-well plates</td><td>Corning incorporated</td><td>CLS3516</td><td></td></tr><tr><td>Spare Chopping Discs&amp;nbsp;</td><td>TED PELLA</td><td>Model 10180-01</td><td></td></tr><tr><td>Tissue chopper</td><td>McIlwain</td><td>Model TC752</td><td>10180-220</td></tr><tr><td>Tweezers, pointed</td><td></td><td></td><td></td></tr><tr><td>Urinary cups</td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Antibodies&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Alexa fluor 488 donkey anti Rabbit IgG</td><td>&amp;nbsp;&lt;br/&gt; Invitrogen</td><td>A21206</td><td>1:500; secondary ab; Secondary dilution buffer;&amp;nbsp; Incubation for 2 h at room temperature</td></tr><tr><td>Anti glial fibrillary acidic protien</td><td>Milliphore</td><td>MAB3402</td><td>1:1500; primary ab; blocking; Host:mouse</td></tr><tr><td>Anti-calbindin-D-28K</td><td>Swant&amp;nbsp;</td><td>CB300</td><td>1:2000; primary ab; blocking; host: rabbit&amp;nbsp;</td></tr><tr><td>Anti-calbindin-D-28K</td><td>Swant</td><td>CB-38a</td><td>1:2000; primary ab; blocking&amp;nbsp;</td></tr><tr><td>Anti-pan-ADP-ribose reagent</td><td>Milliphore</td><td>MABE1016</td><td>1:800; primary ab; blocking; incubation for 2 h at room temperature</td></tr><tr><td>DAPI (4&#039;,6-Diamidine-2&#039;-phenylindole dihydrochloride)</td><td>Cell signaling</td><td>4084</td><td>1:1000; secondary dilution buffer; incubation for 20 min at room temperature</td></tr></tbody></table>

visualizing the dna damage response in purkinje cells using cerebellar organotypic cultures

Cerebellar Purkinje cells (PCs) exhibit a unique interplay of high metabolic rates, specific chromatin architecture, and extensive transcriptional activity, making them particularly vulnerable to DNA damage. This necessitates an efficient DNA damage response (DDR) to prevent cerebellar degeneration, often initiated by PC dysfunction or loss. A notable example is the genome instability syndrome, ataxia-telangiectasia (A-T), marked by progressive PC depletion and cerebellar deterioration. Investigating DDR mechanisms in PCs is vital for elucidating the pathways leading to their degeneration in such disorders. However, the complexity of isolating and cultivating PCs in vitro has long hindered research efforts. Murine cerebellar organotypic (slice) cultures offer a feasible alternative, closely mimicking the in vivo tissue environment. Yet, this model is constrained to DDR indicators amenable to microscopic imaging. We have refined the organotypic culture protocol, demonstrating that fluorescent imaging of protein-bound poly(ADP-ribose) (PAR) chains, a rapid and early DDR indicator, effectively reveals DDR dynamics in PCs within these cultures, in response to genotoxic stress.

Figure 1&#160;illustrates the general appearance of the cerebellar organotypic cultures. The top row in Figure 1A shows the cerebellar foliation, which is maintained in culture, while the bottom row shows the PCs stained for Calbindin D-28k (green) and the neuronal nuclei stained for NeuN (red). In Figure 1B, the astrocytes in the PCs (red) are stained for GFAP (green), an intermediate filament type III protein. <strong class="xfig"...

Author Spotlight: Deciphering the Role of ATM in Ataxia-Telangiectasia and the Associated Cerebellar Degeneration

Cerebellar Purkinje cells (PCs) are particularly sensitive to deficiencies in the DNA damage response. A protocol is presented for the visual evaluation of the dynamics of PC response to DNA damage, which involves staining protein-bound poly(ADP-ribose) chains within cerebellar organotypic cultures.

Visualizing the DNA Damage Response in Purkinje Cells Using Cerebellar Organotypic Cultures

Cerebellar Purkinje cells (PCs) exhibit a unique interplay of high metabolic rates, specific chromatin architecture, and extensive transcriptional ...

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1.4K Views.  Tel Aviv University. Cerebellar Purkinje cells (PCs) are particularly sensitive to deficiencies in the DNA damage response. A protocol is presented for the visual evaluation of the dynamics of PC response to DNA damage, which involves staining protein-bound poly(ADP-ribose) chains within cerebellar organotypic cultures.

Video: Author Spotlight: Deciphering the Role of ATM in Ataxia-Telangiectasia and the Associated Cerebellar Degeneration

Organotypic Cerebellar Cultures: Apoptotic Challenges and Detection

Organotypic cultures of neuronal tissue were first introduced by Hogue in 1947 1,2 and have constituted a major breakthrough in the field of neuroscience. Since then, the technique was developed further and currently there are many different ways to prepare organotypic cultures. The method presented here was adapted from the one described by Stoppini et al. for the preparation of the slices and from Gogolla et al. for the staining procedure 3,4.
A unique feature of this technique is that it allows you to study different parts of the brain such as hippocampus or cerebellum in their original structure, providing a big advantage over dissociated cultures in which all the cellular organization and neuronal networks are disrupted. In the case of the cerebellum it is even more advantageous because it allows the study of Purkinje cells, extremely difficult to obtain as dissociated primary culture. This method can be used to study certain developmental features of the cerebellum in vitro, as well as for electrophysiological and pharmacological experiments in both wild type and mutant mice.
The method described here was designed to study the effect of apoptotic stimuli such as Fas ligand in the developing cerebellum, using TUNEL staining to measure apoptotic cell death. If TUNEL staining is combined with cell type specific markers, such as Calbindin for Purkinje cells, it is possible to evaluate cell death in a cell population specific manner. The Calbindin staining also serves the purpose of evaluating the quality of the cerebellar cultures.

This method describes the generation of organotypic cerebellar cultures and the effect of certain apoptotic stimuli on the viability of different cerebellar cell types.

Organotypic cultures of neuronal tissue were first introduced by Hogue in 1947 1,2 and have constituted a major breakthrough in the field of neuroscience. ...

Neuroscience

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 ...

Visualizing the Beating Heart in Drosophila

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in myogenic hearts. A key step in examining heart function in the fly is finding a way to access the heart in a manner that preserves its myogenic function while still allowing the beating heart organ to be observed and recorded. Two different methods for observing and recording the beating heart in both larva and adult Drosophila are described here. Our semi-intact preparation using adult flies allows clear visualization of the abdominal heart without interference from the pigmented cuticle and overlying fat bodies. To record larval heart beats it is necessary to immobilize the larva, which minimizes body wall movements thereby reducing heart movements that are not associated with myocardial contractions. Our methodologies produce stable adult and larval heart preparations that can beat for hours at rates of 1-3 Hz.

Visualizing the Beating Heart in Drosophila

Technique required for visualizing the beating heart in larval and adult Drosophila are presented. Each life stage requires a different methodology.

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in ...

Quantification of &gamma;H2AX Foci in Response to Ionising Radiation

DNA double-strand breaks (DSBs), which are induced by either endogenous metabolic processes or by exogenous sources, are one of the most critical DNA lesions with respect to survival and preservation of genomic integrity. An early response to the induction of DSBs is phosphorylation of the H2A histone variant, H2AX, at the serine-139 residue, in the highly conserved C-terminal SQEY motif, forming &gamma;H2AX1. Following induction of DSBs, H2AX is rapidly phosphorylated by the phosphatidyl-inosito 3-kinase (PIKK) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)2. Typically, only a few base-pairs (bp) are implicated in a DSB, however, there is significant signal amplification, given the importance of chromatin modifications in DNA damage signalling and repair. Phosphorylation of H2AX mediated predominantly by ATM spreads to adjacent areas of chromatin, affecting approximately 0.03% of total cellular H2AX per DSB2,3. This corresponds to phosphorylation of approximately 2000 H2AX molecules spanning ~2 Mbp regions of chromatin surrounding the site of the DSB and results in the formation of discrete &gamma;H2AX foci which can be easily visualized and quantitated by immunofluorescence microscopy2. The loss of &gamma;H2AX at DSB reflects repair, however, there is some controversy as to what defines complete repair of DSBs; it has been proposed that rejoining of both strands of DNA is adequate however, it has also been suggested that re-instatement of the original chromatin state of compaction is necessary4-8. The disappearence of &gamma;H2AX involves at least in part, dephosphorylation by phosphatases, phosphatase 2A and phosphatase 4C5,6. Further, removal of &gamma;H2AX by redistribution involving histone exchange with H2A.Z has been implicated7,8. Importantly, the quantitative analysis of &gamma;H2AX foci has led to a wide range of applications in medical and nuclear research. Here, we demonstrate the most commonly used immunofluorescence method for evaluation of initial DNA damage by detection and quantitation of &gamma;H2AX foci in &gamma;-irradiated adherent human keratinocytes9.

Quantification of &#947;H2AX Foci in Response to Ionising Radiation

Quantification of DNA double-strand streaks using &gamma;H2AX formation as a molecular marker has become an invaluable tool in radiation biology. Here we demonstrate the use of an immunofluorescence assay for quantification of &gamma;H2AX foci after exposure of cells to radiation.

DNA double-strand breaks (DSBs), which are induced by either endogenous metabolic processes or by exogenous sources, are one of the most critical DNA ...

Modeling Neuronal Death and Degeneration in Mouse Primary Cerebellar Granule Neurons

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their post-natal development, abundance, and accessibility, CGNs are an ideal model to study neuronal processes, including neuronal development, neuronal migration, and physiological neuronal activity stimulation. In addition, CGN cultures provide an excellent model for studying different modes of cell death including excitotoxicity and apoptosis. Within a week in culture, CGNs express N-methyl-D-aspartate (NMDA) receptors, a specific ionotropic glutamate receptor with many critical functions in neuronal health and disease. The addition of low concentrations of NMDA in conjunction with membrane depolarization to rodent primary CGN cultures has been used to model physiological neuronal activity stimulation while the addition of high concentrations of NMDA can be employed to model excitotoxic neuronal injury. Here, a method of isolation and culturing of CGNs from 6 day old pups as well as genetic manipulation of CGNs by adenoviruses and lentiviruses are described. We also present optimized protocols on how to stimulate NMDA-induced excitotoxicity, low-potassium-induced apoptosis, oxidative stress and DNA damage following transduction of these neurons.

This protocol describes a simple method for isolating and culturing primary mouse cerebral granule neurons (CGNs) from 6-7 day old pups, efficient transduction of CGNs for loss and gain of function studies, and modelling NMDA-induced neuronal excitotoxicity, low-potassium-induced cell death, DNA-damage, and oxidative stress using the same culture model.

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their ...

Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate mutations and chromosomal aberrations1. To prevent this trauma from catalyzing genomic instability, it is crucial for cells to detect DSBs, activate the DNA damage response (DDR), and repair the DNA. When stimulated, the DDR works to preserve genomic integrity by triggering cell cycle arrest to allow for repair to take place or force the cell to undergo apoptosis. The predominant mechanisms of DSB repair occur through nonhomologous end-joining (NHEJ) and homologous recombination repair (HRR) (reviewed in2). There are many proteins whose activities must be precisely orchestrated for the DDR to function properly. Herein, we describe a method for 2- and 3-dimensional (D) visualization of one of these proteins, 53BP1.
The p53-binding protein 1 (53BP1) localizes to areas of DSBs by binding to modified histones3,4, forming foci within 5-15 minutes5. The histone modifications and recruitment of 53BP1 and other DDR proteins to DSB sites are believed to facilitate the structural rearrangement of chromatin around areas of damage and contribute to DNA repair6. Beyond direct participation in repair, additional roles have been described for 53BP1 in the DDR, such as regulating an intra-S checkpoint, a G2/M checkpoint, and activating downstream DDR proteins7-9. Recently, it was discovered that 53BP1 does not form foci in response to DNA damage induced during mitosis, instead waiting for cells to enter G1 before localizing to the vicinity of DSBs6. DDR proteins such as 53BP1 have been found to associate with mitotic structures (such as kinetochores) during the progression through mitosis10.
In this protocol we describe the use of 2- and 3-D live cell imaging to visualize the formation of 53BP1 foci in response to the DNA damaging agent camptothecin (CPT), as well as 53BP1's behavior during mitosis. Camptothecin is a topoisomerase I inhibitor that primarily causes DSBs during DNA replication. To accomplish this, we used a previously described 53BP1-mCherry fluorescent fusion protein construct consisting of a 53BP1 protein domain able to bind DSBs11. In addition, we used a histone H2B-GFP fluorescent fusion protein construct able to monitor chromatin dynamics throughout the cell cycle but in particular during mitosis12. Live cell imaging in multiple dimensions is an excellent tool to deepen our understanding of the function of DDR proteins in eukaryotic cells.

This protocol describes a method for visualizing a DNA double-strand break signaling protein activated in response to DNA damage as well as its localization during mitosis.

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate ...

Visualizing Bacteria in Nematodes using Fluorescent Microscopy

Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on earth, nematodes and bacteria form a wide array of symbiotic associations that range from beneficial to pathogenic 1-3. One such association is the mutually beneficial relationship between Xenorhabdus bacteria and Steinernema nematodes, which has emerged as a model system of symbiosis 4. Steinernema nematodes are entomopathogenic, using their bacterial symbiont to kill insects 5. For transmission between insect hosts, the bacteria colonize the intestine of the nematode's infective juvenile stage 6-8. Recently, several other nematode species have been shown to utilize bacteria to kill insects 9-13, and investigations have begun examining the interactions between the nematodes and bacteria in these systems 9.
We describe a method for visualization of a bacterial symbiont within or on a nematode host, taking advantage of the optical transparency of nematodes when viewed by microscopy. The bacteria are engineered to express a fluorescent protein, allowing their visualization by fluorescence microscopy. Many plasmids are available that carry genes encoding proteins that fluoresce at different wavelengths (i.e. green or red), and conjugation of plasmids from a donor Escherichia coli strain into a recipient bacterial symbiont is successful for a broad range of bacteria. The methods described were developed to investigate the association between Steinernema carpocapsae and Xenorhabdus nematophila 14. Similar methods have been used to investigate other nematode-bacterium associations 9,15-18and the approach therefore is generally applicable.
The method allows characterization of bacterial presence and localization within nematodes at different stages of development, providing insights into the nature of the association and the process of colonization 14,16,19. Microscopic analysis reveals both colonization frequency within a population and localization of bacteria to host tissues 14,16,19-21. This is an advantage over other methods of monitoring bacteria within nematode populations, such as sonication 22or grinding 23, which can provide average levels of colonization, but may not, for example, discriminate populations with a high frequency of low symbiont loads from populations with a low frequency of high symbiont loads. Discriminating the frequency and load of colonizing bacteria can be especially important when screening or characterizing bacterial mutants for colonization phenotypes 21,24. Indeed, fluorescence microscopy has been used in high throughput screening of bacterial mutants for defects in colonization 17,18, and is less laborious than other methods, including sonication 22,25-27and individual nematode dissection 28,29.

To study the mutualism between Xenorhabdus bacteria and Steinernema nematodes, methods were developed to monitor bacterial presence and location within nematodes. The experimental approach, which can be applied to other systems, entails engineering bacteria to express the green fluorescent protein and visualizing, using fluorescence microscopy bacteria within the transparent nematode.

 
Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on ...

Preparation and Immunostaining of Myelinating Organotypic Cerebellar Slice Cultures

In the nervous system, myelin is a complex membrane structure generated by myelinating glial cells, which ensheathes axons and facilitates fast electrical conduction. Myelin alteration has been shown to occur in various neurological diseases, where it is associated with functional deficits. Here, we provide a detailed description of an ex vivo model consisting of mouse organotypic cerebellar slices, which can be maintained in culture for several weeks and further be labeled to visualize myelin.

Here we present a method to prepare organotypic slice cultures from mouse cerebellum and myelin sheath staining by immunohistochemistry suitable for investigating mechanisms of myelination and remyelination in the central nervous system.

In the nervous system, myelin is a complex membrane structure generated by myelinating glial cells, which ensheathes axons and facilitates fast electrical ...

Purkinje Cell Survival in Organotypic Cerebellar Slice Cultures

Organotypic slice culture is a powerful in vitro model that mimicks in vivo conditions more closely than dissociated primary cell cultures. In early postnatal development, cerebellar Purkinje cells are known to go through a vulnerable period, during which they undergo programmed cell death. Here, we provide a detailed protocol to perform mouse organotypic cerebellar slice culture during this critical time. The slices are further labeled to assess Purkinje cell survival and the efficacy of neuroprotective treatments. This method can be extremely valuable to screen for new neuroactive molecules.

Organotypic slice cultures are a powerful tool to study neurodevelopmental or degenerative/regenerative processes. Here, we describe a protocol that models the neurodevelopmental death of Purkinje cells in mouse cerebellar slice cultures. This method may benefit research in neuroprotective drug discovery.

Organotypic slice culture is a powerful in vitro model that mimicks in vivo conditions more closely than dissociated primary cell cultures. In early ...

Analyzing Purkinje Cell Survival in Organotypic Cerebellar Slice Cultures

Source: Rakotomamonjy, J., et al. <a href="https://app.jove.com/v/60353">Purkinje Cell Survival in Organotypic Cerebellar Slice Cultures.</a> J. Vis. Exp. (2019)
This video illustrates a technique for obtaining cerebellar organotypic slices from a mouse pup&#39;s brain and culturing the slices in media supplemented with nutrients and growth factors to mimic cerebellar development in vitro. Additionally, specific cerebellar slices are treated with a test compound, and the compound&#39;s neuroprotective ability is analyzed using an immunofluorescence staining technique.

1. Preparation prior to organotypic cerebellar slice cultures

	In a cell culture hood sprayed with 70% ethanol beforehand, prepare 200 mL of culture ...

Visualizing the DNA Damage Response in Purkinje Cells Using Cerebellar Organotypic Cultures

Summary

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Chapters in this video

Organotypic Cerebellar Cultures: Apoptotic Challenges and Detection

The Analysis of Purkinje Cell Dendritic Morphology in Organotypic Slice Cultures

Visualizing the Beating Heart in Drosophila

Quantification of γH2AX Foci in Response to Ionising Radiation

Modeling Neuronal Death and Degeneration in Mouse Primary Cerebellar Granule Neurons

Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins

Visualizing Bacteria in Nematodes using Fluorescent Microscopy

Preparation and Immunostaining of Myelinating Organotypic Cerebellar Slice Cultures

Purkinje Cell Survival in Organotypic Cerebellar Slice Cultures

Analyzing Purkinje Cell Survival in Organotypic Cerebellar Slice Cultures