Excitotoxic necrosis is a leading form of neurodegeneration. This process of regulated necrosis is triggered by the synaptic accumulation of the neurotransmitter glutamate, and the excessive stimulation of its postsynaptic receptors. However, information on the subsequent molecular events that culminate in the distinct neuronal swelling morphology of this type of neurodegeneration is lacking. Other aspects, such as changes in specific subcellular compartments, or the basis for the differential cellular vulnerability of distinct neuronal subtypes, remain under-explored. Furthermore, a range of factors that come into play in studies that use in vitro or ex vivo preparations might modify and distort the natural progression of this form of neurodegeneration. It is therefore important to study excitotoxic necrosis in live animals by monitoring the effects of interventions that regulate the extent of neuronal necrosis in the genetically amenable and transparent model system of the nematode Caenorhabditis elegans. This protocol describes methods of studying excitotoxic necrosis in C. elegans neurons, combining optical, genetic, and molecular analysis. To induce excitotoxic conditions in C. elegans, a knockout of a glutamate transporter gene (glt-3) is combined with a neuronal sensitizing genetic background (nuls5 [Pglr-1::GαS(Q227L)]) to produce glutamate receptor hyperstimulation and neurodegeneration. Nomarski differential interference contrast (DIC), fluorescent, and confocal microscopy in live animals are methods used to quantify neurodegeneration, follow subcellular localization of fluorescently labeled proteins, and quantify mitochondrial morphology in the degenerating neurons. Neuronal Fluorescence Activated Cell Sorting (FACS) is used to distinctly sort at-risk neurons for cell-type specific transcriptomic analysis of neurodegeneration. A combination of live imaging and FACS methods as well as the benefits of the C. elegans model organism allow researchers to leverage this system to obtain reproducible data with a large sample size. Insights from these assays could translate to novel targets for therapeutic intervention in neurodegenerative diseases.
Excitotoxicity is the leading cause of neuronal death in brain ischemia and a contributing factor in multiple neurodegenerative diseases1,2,3,4,5,6,7,8,9. Disruption of oxygenated blood flow to the brain (e.g., due to a blood clot) results in the malfunction of glutamate transporters, leading to accumulation of glutamate in the synapse. This excess of glutamate over-activates post-synaptic Glutamate Receptors (GluRs) leading to an excessive (catalytic, non-stoichiometric) influx of Ca2+ into neurons (Figure 1A). This detrimental influx leads to progressive postsynaptic neurodegeneration that morphologically and mechanistically ranges from apoptosis to regulated necrosis10,11,12. Although they were based on successful interventions in animal models, multiple clinical trials of GluR antagonists that sought to block Ca2+ entry and promote cell viability have failed in the clinical setting13,14,15,16. A likely critical contributor to these failures is the fact that (in contrast to the animal models) treatment in the clinical setting is administered hours after stroke onset, causing the intervention to block late-acting neuroprotective mechanisms, while failing to interrupt degenerative signaling downstream of GluRs14,16,17. An alternative approach, which is based on thrombolysis, can only be administered within a severely restricted time window, leaving many patients (who suffer stroke at home with poorly identifiable time of onset) unable to benefit from it17. These setbacks emphasize the need to focus excitotoxicity research on the study of events occurring after GluR hyper-stimulation and differentiate subsequent degenerative cascades from concurrent neuroprotective processes. This approach can help prevent cell damage and identify efficient drug targets that can be administered later after damage onset.
One approach to identify subsequent events in excitotoxicity is to study the cell-death signaling mechanisms downstream of GluR hyperstimulation, such as those leading to mitochondrial collapse. Drastic malfunction of mitochondrial physiology and dynamics is a hallmark of neurodegeneration, as seen in excitotoxicity18,19,20. While all cells depend on mitochondrial function and availability for survival, activity, and cellular maintenance, neurons are particularly dependent on mitochondrial energy production to support signal transmission and propagation. Specifically, neurons spend ~50% of their signaling-related energy consumption to restore resting membrane potential following the activation of postsynaptic receptors/channels21, with high dependence on oxygen and glucose. The reduced availability of glucose and oxygen observed in stroke leads to serious mitochondrial alterations, causing further reduction in ATP production19,22,23,24. However, studies to identify the sequence of events that lead to mitochondrial collapse produced controversial results and lacked consensus. Analyzing mitochondrial morphology can help understand these events leading to mitochondrial pathology since it is a good indicator of neuronal health25,26,27,28,29. Filamentous mitochondria are representative of a healthy neuron, whereas fragmented mitochondria reveal substantial neuronal damage that could lead to cell death. Analyzing mitochondrial morphology in live animals under different genetic conditions can help focus on specific genes and pathways involved in mitochondrial-dependent neurodegeneration in excitotoxicity.
Another approach to identifying subsequent events that might regulate the extent of excitotoxic neurodegeneration is to study the transcriptional neuroprotective mechanisms that mitigate some of the effects of excitotoxicity14,16. However, the lack of specificity of key neuroprotective transcription factors and the divergence of experimental setups impede the success of efforts to clearly identify core neuroprotective programs (especially in regulated necrosis).
Therefore, both the study of downstream death signaling pathways and the study of transcriptional neuroprotection in excitotoxicity encountered great difficulties and disagreements on observed outcomes. Much of this controversy is likely to arise from the use of ex vivo or in vitro models of excitotoxicity, and the variability introduced by the specificity of different experimental setups. It is therefore highly beneficial to focus on identifying core mechanisms that are highly conserved, and study them in vivo. The simple model system of the nematode C. elegans offers a particularly effective option, due to the potent combination of particularly powerful and diversified research tools, the conservation of core cell-death pathways, and the rich information on the structure and connectivity of its nervous system30,31,32,33. Indeed, the seminal work of the Driscoll lab on the genetic analysis of necrotic neurodegeneration in mechanosensory neurons is an excellent demonstration of the power of this approach34. Importantly for the analysis of excitotoxicity, the conservation of signaling pathways in the nematode includes all major components of glutamatergic neurotransmission35,36.
The nematode excitotoxicity model builds on these seminal studies, allowing the researcher to study biochemical processes akin to those that occur in stroke and other neurodegenerative diseases affected by glutamate-dependent neurotoxicity. To induce excitotoxic conditions in C. elegans this experimental approach uses an excitotoxicity strain that is the combination of a knockout of a glutamate transporter gene (glt-3) and neuronal sensitizing genetic background (nuls5 [Pglr-1::GαS(Q227L)]) to produce GluR hyperstimulation and neurodegeneration37. This excitotoxicity strain exposes 30 specific (glr-1-expressing) neurons that are postsynaptic to glutamatergic connections to excitotoxic neurodegeneration. Of these 30 at-risk neurons, individual neurons go through necrosis as the animal progresses through development (with mixed stochasticity and partial preference towards certain specific neurons38), while at the same time cell corpses are also being gradually removed. In combination with the accessibility of many mutant strains, this approach allows the study of multiple pathways that affect neurodegeneration and neuroprotection. These approaches have already been used to analyze some of the downstream death signaling cascades39 and transcriptional regulators of excitotoxic neurodegeneration in excitotoxicity38,40. Like other cases of necrotic neurodegeneration in the worm41, the nematode's excitotoxic neurodegeneration does not involve classic apoptosis40.
This methods paper describes the basic system to induce, quantify, and manipulate excitotoxic necrotic neurodegeneration in C. elegans. Furthermore, it outlines two main protocols that are currently in use to streamline studies of specific aspects of nematode excitotoxicity. By using fluorescent reporters and live in vivo imaging the researcher can study mitochondrial involvement and dynamics in the nematode model of excitotoxic neurodegeneration. To determine the effect of specific neuroprotective transcription factors, the investigator can use cell type-specific expression of fluorescent markers, dissociation of animals into single cells, and FACS to isolate specific neurons that are at risk of necrosis from excitotoxicity. These cell-type specific isolated neurons can then be used for RNA sequencing in strains that harbor mutations in key transcription factors. Put together, these methods can allow researchers to tease out the molecular underpinnings of excitotoxic neurodegeneration and neuroprotection in vivo with great clarity and precision.
1. Strains used to Investigate Excitotoxic Neurodegeneration & Neuroprotection
- Use the nematode excitotoxicity strain ZB1102 as the reference point for standard excitotoxic neurodegeneration.
NOTE: Glutamate-dependent excitotoxicity in C. elegans is produced in strain ZB1102 by combining a knockout (ko) of a glutamate transporter with a transgene that sensitizes neurons in these animals to neurotoxicity and is expressed in a subset of neurons postsynaptic to glutamatergic connections37. This genetic combination is referred to as the nematode excitotoxicity strain, and it is freely available from the Caenorhabditis Genetics Center (CGC).
- To study the effect of genes encoding candidate regulators of excitotoxic necrosis, combine a mutation in such genes (e.g., dapk-1, or crh-1) with the nematode excitotoxicity strain.
- Conduct genetic crosses according to standard C. elegans methods30, as outlined in wormbook.org42,43.
- Since the molecular basis of the mutation is typically documented, follow the cross progeny by genotyping the specific locus using PCR. Differentiate WT vs mutant by fragment size (for deletions) or sequencing (for point mutations).
NOTE: Table 1 outlines strains that were used to study distinct pathways of neurodegeneration and neuroprotection, specifically for this protocol.
- Derive all critical strains by two independent lines and test them separately to confirm the validity of the observed phenotype.
2. Growth Media and Animal Husbandry
- Grow worms at 16-25 °C on either standard NGM plates30,37,42 or MYOB plates38,44 seeded with OP50 E.coli as per standard methods.
NOTE: MYOB plates give identical results as NGM plates but are slightly easier to prepare.
- Maintain the experimental strains consistently well fed.
NOTE: Starvation can affect neurodegeneration and reduce the number of dying head neurons. If worms are poorly fed or starved, plate them onto fresh plates and wait a few generations before continuing experiments. Transgenerational effect of starvation will wane after a few generations.
3. Quantification of degenerating head neurons by Nomarski differential interference contrast (DIC) and scoring
- Use the nematode excitotoxicity strain (ZB1102: glt-3(bz34) IV; nuIs5 V) as the experimental control for quantification, representing normal excitotoxicity levels. The protocol does not follow the same animal through different developmental stages. Instead, it gives a snapshot of a mixed-stage population of animals, so that in total one collects information from different animals to represent all developmental stages.
NOTE: The nuls5 transgene[Pglr-1::GαS(Q227L). Pglr-1::GFP]45 (which expresses an activated Gαs and GFP in neurons postsynaptic to glutamatergic connections) produces a background GluR-independent necrotic neurodegeneration level of ~1 dying head neuron/animal (at any given time during development). The addition of glt-3 (ko) increases necrotic neurodegeneration of the postsynaptic nuIs5-expressing neurons in a GluR-dependent manner, going up to 4-5 dying head neurons/animal in the third larval stage (L3)37.
- For the test strains, use animals from a recently completed genetic cross, or thaw strains of interest from -80 °C frozen stocks prepared from fresh crosses. Wait a few generations (≥4) before scoring a phenotype and neurodegeneration.
- Cut and remove a small piece of agar from a plate of a mixed-stage population of well-fed animals, and mount it onto a coverslip by flipping the agar chunk over, so that the animals that were crawling on the surface of the agar now face the coverslip.
NOTE: Animals can still move, but are now somewhat restrained and can be observed without anesthetic. Such a chunk can be used for ~1 hour before replacing it with a fresh one.
- Using an inverted DIC scope with x10 ocular and either x40 or x63 objective, scan animals at random by sliding the coverslip manually. While other imaging steps described below can be conducted on either inverted or upright microscopes, the examination of worms in the agar chunk requires an inverted scope.
- Identify the developmental stage of each animals by the shape of its uterus (refer to uterus diagrams in wormbook.org).
- For each animal, record its developmental stage and the number of dying (i.e., vacuolated) neurons. Count and record the total number of dying neurons in the head, the number of dying neurons in the retrovesicular ganglion (which gives easy identification of specific cells), and the number of dying neurons in the tail (which is unaffected by glutamate and serves as an internal control, confirming that the sensitizing transgene nuIs5 is fully active).
- Record this data in a table where animals from a given strain are grouped by categories of developmental stage.
- In each session, collect data at random from worms of several developmental stages; perform several scoring sessions across multiple days to collect enough data for statistical analysis. Record neurodegeneration levels at multiple stages and construct a bar graph similar to Figure 1C.
NOTE: This method will allow one to determine if a certain treatment or mutation raises/decreases neurodegeneration levels at all stages (shifting the distribution up/down) or shifts the peak in neurodegeneration to an earlier or later developmental stage (shifting the distribution left/right).
- Perform data collection while blinded to the identity of the genotype. Confirm in two independent isolates of the genetic line (to minimize the danger of unintended effects of other/unexpected genetic differences between isolates), and pool data for analysis.
- In each strain, calculate the average and SEM of the number of degenerating neurons in the head (including the retrovesicular ganglion) in each developmental stage (Figure 1D). Perform a similar analysis of dying retrovesicular ganglion -only and tail neurons, as necessary.
4. Identification of specific degenerating head neurons
- Mount individual (or small groups of-) animals on an agar pad46, and paralyze the worm with tetramisole (see below, section 5).
- With a combined fluorescence and DIC scope (upright or inverted) locate specific vacuolated neurons.
- Determine the specific neuron identity of the vacuolated neuron by tracking its GFP labeled processes and comparing the location of the cell body and shape of the processes to those of the neurons known to expresses glr-1 using WormAtlas47.
NOTE: Alternatively, identification can be assisted by new methods to quickly identify neurons by multi-color labeling that are coming soon from the Hobert lab48.
5. Live imaging of neuronal mitochondrial morphology by fluorescent microscopy of reporter strains
- For imaging of mitochondrial changes in degenerating postsynaptic neurons (which are labeled with cytoplasmic GFP in the original excitotoxicity strain) examine mito-mCherry fluorescence.
- Cross the test excitotoxicity strain with strains that label mitochondria of postsynaptic neurons with red fluorescence (by expressing a fusion between the fluorescent protein and the N-terminal of TOM-20 under glr-1 promoter; for details on constructs and strains, see49). Animals can now be imaged using a regular epifluorescence microscope (upright or inverted) or a confocal microscope.
- To paralyze the worm without affecting neuronal survival, pipette 5 µL of 10 mM tetramisole onto a freshly prepared agarose pad. See Arnold et al.46 for detailed protocol on pad preparation.
- Place animals onto the center of the tetramisole drop and mount with a coverslip.
- Seal the sides of the coverslip with nail polish and allow the nail polish to dry.
- Within 20 minutes of tetramisole treatment and using a scope with DIC and fluorescence imaging, locate worms using a 20x objective lens.
- Once the head of the worm is located (or neurons of interest) move to a 100x oil objective and focus on the fluorescence labeling of the mitochondria in the soma.
- Capture Z-stack images using DIC, GFP, and TxRed filter settings.
6. Neuronal mitochondria morphology scoring & quantification
- Analyze mitochondrial morphology either during live imaging of the worm or after image acquisition using ImageJ or other imaging software.
- For easy identification and repeated analysis of specific neurons, identify the three neurons in the retrovesicular ganglion, RIGL/R and AVG (in some cases only two are present due to cell corpse clearance in excitotoxicity).
- Categorize mitochondria into three main groups: filamentous, intermediate, and fragmented50,51,52.
NOTE: Filamentous mitochondria appear as continuous thin structures in the soma of neurons, usually surrounding the nucleus. Intermediate mitochondria appear as a combination of at least one apparent filamentous network, albeit with breaks, and some fragmentation in the soma.Fragmented mitochondria exhibit complete breaks in the mitochondrial network, have swollen appearance, and are scattered throughout the soma (Figure 2A).
- For each worm, score the percent of neurons with filamentous, intermediate, or fragmented mitochondria for a total of at least 30 worms.
- Perform statistical analysis using one-way ANOVA followed by post hoc Tukey's test for each mitochondrial morphology between strains53.
7. Buffer and reagent preparation for worm dissociation for FACS of neurons at risk of neurodegeneration
- Prepare M9 buffer as follows: 3 g of KH2PO4, 6 g of Na2HPO4, 5 g of NaCl, H2O to 1 L. Sterilize by autoclaving. Add 1 mL of filter-sterilized 1 M MgSO4. Store at room temperature.
- Prepare Egg buffer as follows: 118 mM NaCl, 48 mM KCl, 2 mM CaCl2, 2 mM MgCl2 ; autoclave to sterilize. Add 2 M of HEPES pH 7.3 stock solution (previously filtered with a 0.2 μm bottle top filter) to a final concentration of 25 mM. Adjust pH to 7.3 with 1 N NaOH (no more than 10 mL). Use osmometer to ensure the final osmolarity is between 335 -345 mOsm. Filter sterilize egg buffer with 0.2 μm bottle top filters. Store in 4 °C.
NOTE: The salts dissolve easier when using MgCl2 ·6H2O and CaCl2·2H2O to make respective MgCl2 and CaCl2 stock solutions. You can also make a stock of 10x egg buffer and dilute it when needed in sterile deionized water.
- Prepare SDS-DTT as follows: 20 mM HEPES pH 8.0, 0.25% SDS, 200 mM DTT, 3% sucrose. In a tissue culture hood, sterilize SDS-DTT with 0.2 μm syringe filter. Store 300 µL aliquots in -20 °C covered in foil to protect from light.
- Prepare pronase solution as follows: prepare the day of the dissociation 15 mg/mL pronase in egg buffer. Store on ice.
8. Age Synchronization for neuron specific FACS
- Use animals that combine the excitotoxicity genotype (and other mutations, as desired) with transgenic expression of a strong fluorescent marker that can be readily used for sorting (e.g., FJ1244: pzIs29 [Pglr-1::NLS::LAC-Z::GFP::glr-1 3'UTR] X)54. Grow animals on two or three 100mm NGM/MYOB plates at 20 °C until the plate is full of predominantly gravid worms.
- Wash worms off the plates using M9 buffer. Transfer worms into 50 mL conical tubes using a glass 10 mL serological pipette.
- Centrifuge at room temperature for 2.5 min at 250x g and remove the supernatant.
- Resuspend the worm pellet in 10 mL of bleach solution (2% 10 N NaOH and 5% fresh household bleach in sterile deionized water).
- Place tubes on a shaker horizontally, rock at low speed. Ensure that the worms do not settle to the bottom of the tube. The bleach cracks open the worm's cuticle but does not affect embryos, which are protected by the egg shell.
- This bleaching step takes approximately 5 minutes, but this varies. To ensure over-bleaching does not occur, monitor the process by retrieving a sample every minute: Gently pipette 10 µL from each tube on a glass microscope slide and examine the worms using a dissecting microscope.
- Once most of the gravid worms are cracked open but not completely dissolved, stop the bleaching step by filling the conical tubes with egg buffer.
- To retrieve the eggs/embryos, centrifuge the tubes at 250x g for 2.5 min, remove the supernatant and wash again with egg buffer.
- Repeat four more washes.
- Resuspend the eggs by gently pipetting the pellet and spread on four 100 mm seeded NGM/MYOB plates. Let embryos hatch and grow at 20 °C for 3 days.
NOTE: The plates should now be full of predominantly gravid worms.
- Repeat the age synchronization by repeating this bleach/age synchronization protocol.
- Pipette the eggs onto eight 100 mm NGM/MYOB plates. Let animal hatch and grow at 20 °C until at the desired larval stage.
9. Whole worm cell dissociation for FACS
- Grow synchronized worms to the desired developmental stage. Gently wash off 100 mm plates with M9 buffer and glass serological pipettes and transfer to 50 mL conical tubes.
- Add cold M9 up to 45 mL. Place the tubes on ice for 30 minutes to allow the worms to settle at the bottom by gravity, while any residual bacterial debris from the plates will float in the supernatant.
- Remove the supernatant and wash the worms with fresh M9.
- Repeat the 30 minute gravity settling on ice.
- Remove the supernatant, add M9 up to 45mL, and centrifuge at 250 x g for 5 minutes.
- Transfer the pellet to a microcentrifuge tube and add M9 up to 1 mL.
- Spin at 14,000 rpm on a tabletop centrifuge for 1 min and remove supernatant.
- To disrupt the cuticle, resuspend the worm pellet with SDS-DTT using (approximately) twice the volume of the pellet, and incubate with rocking at room temperature for 4 minutes for L2-adult stages.
NOTE: Do not exceed 4 minutes incubation in SDS-DTT because fluorescent protein signal will drastically decrease with longer SDS-DTT treatment. Since the SDS-DTT is light sensitive it should not be more than 3 months old because the solution may have lost potency with time; Dissociations work best with recently -prepared SDS-DTT solution.
- Add 1x egg buffer (pH 7.3, 335-345 mOsm) up to 1 mL to stop the SDS-DTT treatment.
- Spin at 14,000 rpm on a tabletop centrifuge for 1 minute and remove supernatant.
- To further disrupt the cuticle and dissociate the animals' cells, resuspend pellet with room temperature pronase solution, using triple the volume of the pellet.
- Incubate at room temperature for 15-30 min.
- Pipette up and down 40 times every 5 min. with a P-200 or P-1000 pipette, touching the bottom of the tube with the pipette tip.
NOTE: The pressure created by the tip touching the bottom of the tube facilitates worm dissociation. Use a filter tip so the worms do not accidently enter the pipette shaft during rapid pipetting.
- After 15 min, check the progress of the worm dissociation by gently pipetting 5µL on a glass slide and observing the progress under a dissecting microscope. When most (~90%) of the intact worms have burst, the process is complete.
NOTE: Pronase incubation can be extended if many worms remain intact.
- In a cell culture hood, add 1x egg buffer up to 1.5 mL to stop the pronase treatment.
- Transfer to FACS tubes, add 5mL egg buffer, and spin at 800 x g for 5 min.
- Remove supernatant and resuspend in 3 mL pf egg buffer.
- Place a 70 µm cell strainer cap on new FACS tubes, pipette cell suspension on strainer, and spin at 800 x g for 1 min. Collect the efflux cell suspension.
- Place a 5 µm cell strainer cap on new FACS tubes, pipette cell suspension on strainer, and spin at 800 x g for 1 min.
- Add DAPI for a final concentration of 0.5 µg/mL in egg buffer, place on ice with a lid/cover to protect from light.
- Perform FACS as soon as possible. Fluorescent protein signal decreases with time and light exposure, therefore it is important to perform dissociation protocol as quickly as possible and perform FACS immediately after the dissociation is complete.
NOTE: Worm dissociation protocol was adapted from protocols from the Miller lab55,56,57, the Murphy lab58, and the Shaham lab59.
10. FACS machine operation modifications for identification of C. elegans neurons
- Use 4 liters of chilled (4 °C) egg buffer instead of standard sheath fluid when sorting C. elegans neurons.
NOTE: The osmolarity of C. elegans cells is much higher than mammalian cells and standard sheath fluid will burst the neurons. All of the machine settings and calibrations are done after the egg buffer is added, because the viscosity of the egg buffer is different than that of standard sheath fluid. The sorting technician will run the diagnostic beads through the machine to ensure the lasers are working properly.
- When sorted neurons are to be used in subsequent transcriptomics studies, sort a minimum of 100,000 GFP+ cells directly into 800 µL of Trizol-LS + 10 µL of RNase inhibitor.
- Invert cells in Trizol 15 times to mix.
- Snap freeze in a dry ice/ethanol bath and store at -80 °C until ready to extract RNA from all samples for one RNA sequencing experiment.
NOTE: While most sorted cells are collected directly into Trizol for transcriptome analysis, make sure to retrieve a small sample of GFP+ sorted cells (from each strain) collected into egg buffer, for microscopy validation of the efficiency of sorting.
11.FACS gating strategy
- For identifying GFP positive signal use the 488 nm laser with the 530/30 filter and 502 long pass (LP).
- For identifying DAPI positive and negative cells use the 405 nm laser with the 450/50 filter.
- For identifying auto-fluorescent cells that could be mistaken for GFP-positive cells use the 488 nm laser with the 610/20 filter and 595 LP (personal communication, Stanka Semova, Operations Manager, Flow Cytometry Resource Center, Rockefeller University).
- Perform a standard gating strategy and remove events with a large side scatter area (SSC-A) and small forward scatter area (FSC-A), which are likely to represent clumps of cells and debris.
- Isolate foreword scatter singlets and then side scatter singlets.
- Isolate cells with high GFP signal and low autofluorescence signal.
NOTE: Cells high for autofluorescence and lower in GFP are not true GFP positive cells.
- Threshold for GFP+ gate is determined by comparing GFP- cells (i.e. N2) to GFP+ cells55,56.
- Remove any dead cells with a live/dead gate, based on the selective permeability of DAPI to dead cells: Threshold for live dead gate is determined by comparing unstained cells to DAPI stained cells.
NOTE: When sorting multiple samples, flush the stream between samples to ensure there is no cross contamination.
12. Microscopy of sorted neurons to validate the efficiency of FACS gating strategy
- Draw a circle on a glass microscope slide with a liquid repellent pen.
- Pipette 10 µL of sorted cells suspension in egg buffer inside the circle. Place a coverslip over the sample and seal with nail polish around the perimeter of the cover slip.
- After the nail polish has dried, check under upright/inverted fluorescent microscope that the sorted cells are indeed primarily GFP + cells.
NOTE: Perform this check after each sorting session to validate the FACS gating strategy57.
13. RNA Extraction and RNA quality quantification on a Quality Control automated electrophoresis system
NOTE: All RNA work should be executed with extreme care to avoid contamination with RNase (including careful preparation of reagents, consumables, and best RNase-safe practices).
NOTE: Perform all steps where samples contain phenol and/or chloroform in a fume hood.
- Thaw vials of cells in Trizol at room temperature.
- Using a filter tip P200 pipette tip, pipette up and down several times to homogenize the sample.
- Incubate at room temperature for 5 minutes.
- Add 0.2 mL of chloroform per 1 mL of Trizol reagent used for lysis, then securely cap the tube.
- Invert tubes 15 times, and incubate (at 20-25 °C) for 2-3 minutes.
- Centrifuge the sample for 15 minutes at 12,000 × g at 4 °C. The mixture separates into a lower red phenol-chloroform, an interphase, and a colorless upper aqueous phase.
- Exclusively collect the upper aqueous phase containing the RNA and transfer to a new low DNA/RNA bind 1.5 mL tube.
NOTE: In some instances, if the ratio of cells in egg buffer that dropped out of the cell sorter into Trizol is greater than a 1:3 sample:Trizol-LS ratio, the high salt content of the egg buffer can cause the layers to be inverted; if this occurs add more Trizol-LS and repeat the inversion and centrifugation. This can also be avoided if you take note of the increase in sample volume after sorting; if the 1:3 ratio is not maintained, add sufficient trizol before beginning the RNA extraction.
- To further purify the RNA use RNA extraction column chemistry.
- Use the Quality Control automated electrophoresis system to measure the RNA Integrity Number (RIN) and confirm RNA RIN of 8.0 or greater for input into subsequent RNA sequencing experiments.
Nematode model of excitotoxicity and identification of vacuolated degenerating neurons
Data shown here is reproduced from previous publications37,38. To mimic excitotoxic-induced neurodegeneration, a glutamate-transporter gene knockout (glt-3) is combined with a neuronal sensitizing transgenic background (nuls5 [Pglr-1::GαS(Q227L);Pglr-1::GFP)]). The transgenic construct is expressed in neurons expressing GLutamate Receptor 1 (GLR-1, a critical subunit of AMPA receptors) and is tracked via GFP fluorescence expressed in these same neurons. Degenerating neurons are observed using DIC and appear as swollen vacuoles (Figure 1B). The number of dying head neurons is significantly increased when the nuIs5 transgenic construct is combined with glt-3 ko. This number is greatly reduced when all GluRs that contribute to Glu-induced currents in these cells are knocked out, suggesting Glu-dependent neurodegeneration, which is a hallmark of excitotoxicity (Figure 1C). Figure 1D shows the role of the nematode homolog of the transcription factor CREB in nematode excitotoxicity. Knockout of CREB/CRH-1 significantly increases the number of dying head neurons, demonstrating a conserved role in neuroprotection. Table 1 shows the various strains we use in addition to our excitotoxic strain to study processes that can contribute to neurodegeneration in our nematode model.
Visualization and analysis of neuronal mitochondria using fluorescent reporters and in vivo imaging
To observe mitochondrial morphology in excitotoxicity, expression of the transgene odIs12249(a gift from the Rongo Lab), which labels mitochondria in glr-1 -expressing neurons with mito-mCherry fluorescent protein, was combined with the excitotoxicity strain. Using 100x objective of a scope with DIC and fluorescence imaging, mitochondria in the RIGL/R and AVG neurons of the retrovesicular ganglion were examined and categorized into filamentous, intermediate, or fragmented groups (Figure 2A). Degenerating neurons with vacuolated appearance exhibited robust fragmentation of mitochondria (Figure 2B).
Fluorescence activated cell sorting of glr-1 expressing neurons for transcriptomics
Animals from synchronized cultures of control or worms transgenic for pzIs29 (expressing a nuclear GFP marker under the glr-1 promoter)54 were dissociated into cells, and GFP-expressing neurons were collected by FACS. To validate our sorting gates (Figure 3A & Figure 3B), we use microscopy to screen for GFP expression in sorted neurons versus an unsorted C. elegans sample (Figure 3C).
|KP742||nuIs5 [glr-1::gfp; glr-1::Gαs(Q227L) ; lin-15(+)] V||Expresses constitutively active G alpha subunit in glr-1 -expressing neurons. Acts as a neuronal sensitizing background.||45|
|ZB1102||glt-3(bz34) IV; nuIs5 V||Excitotoxicity Strain: Knockout of glutamate transporter 3 (glt-3) combined with sensitizing background nuIs5 to produce excitotoxicity.||37; 38; 39; 40|
|ZB1336||nmr-1 (ak4) II; glr-2 (ak10) III, glr-1 (ky176) III; glt-3(bz34) IV; nuIs5 V||.Combination of the excitotoxicity strain with the knock out of all GluRs mediating ionotropic Glu response in these neurons||37|
|IMN36||crh-1 (tz2)III; glt-3(bz34) IV; nuIs5 V||Combination of the excitotoxicity strain with the knockout of CREB/crh-1.||38|
|odIs122 [Pglr-1::TOM-20(N-terminus)::mCherry, ttx-3::rfp] X||mCherry fused to the N-terminus of mitochondrial protein TOM-20 expressed in glr-1 expressing neurons.||49|
|IMN66||glt-3(bz34) IV; nuIs5 V; odIs122 X||Expresses mitochondrial fluorescent transgene odIs122 in the excitotoxicity strain.||This manuscript|
|N2||C. elegans wild type isolate.||C. elegans var Bristol. Wild type C. elegans strain used as negative control in FACS experiment.||30|
|FJ1244||pzIs29 [Pglr-1::NLS::LAC-Z::GFP::glr-1 3’UTR] X||Expression of bright nuclear GFP (nGFP) in glr-1 -expressing neurons.||54|
|IMN87||glt-3(bz34) IV; nuIs5 V; pzIs29 X||Combination of the excitotoxicity strain with glr-1::nGFP expression for FACS sorting of at-risk neurons.||This manuscript|
Table 1. Strains that used here to study excitotoxicity and the involvement of mitochondria, and to sort at-risk neurons for transcriptomics analysis.
Figure 1. Nematode Excitotoxicity Model. A) Schematic of Excitotoxicity Mechanism. In stroke, lack of energy for neurotransmitter clearance causes a buildup of synaptic Glutamate (Glu), over-stimulation of GluRs, excessive Ca2+ influx and cell death by regulated necrosis or apoptosis. Cells exposed to non-maximal insult intensity show either delayed death or recovery and survival. This later-stage neuroprotection is largely mediated by TFs that trigger defensive transcriptional programs. B) Identification of Necrotic Neurons. Necrotic neurodegeneration is seen in DIC optics as vacuole-looking structures, mostly near the nerve ring of the worm (approximate scale bars of 10µm). Red arrows indicate degenerating cells (only some of which are clearly seen in this focus plane). C) Quantification of excitotoxic neurodegeneration. A deletion of glt-3 combined with the nuIs5 sensitizing background gives rise to GluR -dependent excitotoxicity, seen by significant increases in levels of neurodegeneration. This extended neurodegeneration is reversed by deletion of all GluRs active in these neurons (reproduced from37). D) The effect of CREB/CRH-1 on excitotoxicity (Reproduced from38). Average number of degenerating head neurons per animal in different developmental stages, comparing the original excitotoxicity strain (glt-3;nuIs5) to a similar strain where CREB/CRH-1 is eliminated (crh-1;glt-3;nuIs5). In this and all subsequent bar graphs: Error bars represent SEM of the number of degenerating head neurons per animal. One-way ANOVAs were performed at each life stage. Asterisks represent statistical significance of the difference between the indicated groups, where (when used-) * indicates p≤0.05; ** indicates p≤0.01; *** indicates p≤0.001, n.s. denotes non-significant difference between groups. Please click here to view a larger version of this figure.
Figure 2. Visualization of neuronal mitochondria using fluorescent reporter A) Categorizing mitochondria based on morphology. Images of at-risk neurons labeled with GFP and mito-mCherry (in strain IMN66: glt-3(bz34) IV; nuIs5 V; odIs122 X) using an inverted scope with DIC and fluorescence imaging. Filamentous mitochondria appear as a continuous, circular structure in neuronal soma, while intermediate mitochondria have thin filaments with partial breaks/small amounts of fragmented mitochondria. Fragmented mitochondria have complete breaks with swollen appearance in the neuronal soma. Blue dashed lines indicate the soma of neurons. B) Mitochondria in a degenerating neuron. Vacuolated neurons which indicate neurodegeneration have apparent fragmented and swollen mitochondria in their soma. (Scale bar: 5µm) Please click here to view a larger version of this figure.
Figure 3. Fluorescence activated cell sorting of glr-1 expressing neurons. Forward scatter singlets gate and GFP + gates shown for samples containing A) Neurons expressing nuclear GFP under the glr-1 promoter, or B) wild type cells (devoid of any transgenes expressing fluorescent proteins). C) Images of FACS sorted GFP+ neurons and unsorted cells used to verify the success of sorting. (Scale bar: 10µm) Please click here to view a larger version of this figure.
While the prevalent controversies and failures suggest that excitotoxicity presents an exceptionally hard process to decipher, the analysis of excitotoxicity in the nematode offers a particularly attractive strategy to illuminate conserved neuronal cell death pathways in this critical form of neurodegeneration. The investigator can rely on the rich collection of research tools available in this system, and particularly on the animal's transparency (allowing in vivo analysis) and the large repertoire of viable mutants (available freely from the CGC or prepared in-house by techniques like EMS mutagenesis and CRISPR). Furthermore, research progress benefits from the relative ease of identification of specific neurons that are known to mediate well-described roles in the nematode nervous system. Indeed, one can now identify and quantify degeneration of specific neurons38 and target them for analysis of mitochondrial damage or transcriptomics. This can allow the identification of cell-type specific processes that confer particular vulnerability or resistance.
The powerful molecular tools available in C. elegans research and the genetic tractability of this model organism empower the researcher to dive deeper into the biology of causes of neuronal cell death and protection during excitotoxicity. Moreover, the transparency of the nematode also allows the study of dynamic processes such as mitochondrial morphology changes in an excitotoxicity specific context in live animals. C. elegans' short life cycle and ease of maintenance allow generation of large data sets, in specific neurons, and under diverse conditions (e.g., aging).
There are a few points of special practical significance. The use of sodium azide is a common approach to immobilizing C. elegans for microscopy60. However, sodium azide is an inhibitor of the mitochondrial respiratory chain and causes neurodegeneration61. It is therefore critical to avoid sodium azide, and it is best to quantify the effect of excitotoxicity modifiers in non-treated animals by examining animals on an agar chunk with DIC optics. When necessary, use a muscle-paralyzing agent (such as tetramisole30,62, though neuronal side effects should be considered).
The transgene expressed in nuls5 animals seems to be subjected to gradual suppression of its expression. As a result, after multiple generations, the effect of the sensitized background begins to wane and reduce baseline neurodegeneration. Therefore, the overall excitotoxicity observed in the worm seems to decline with subsequent generations (e.g., as manifested by the decline in degeneration of tail neurons). To avoid this, outcross the excitotoxic strain to wildtype (N2) worms at least twice using standard protocols, and use freshly outcrossed animals, or frozen stock thereof, for scoring of neurodegeneration. Occasional outcrossing restores overall excitotoxicity and neurodegeneration compared to multi-generational lines.
This protocol describes imaging of neuronal mitochondria in live animals during excitotoxicity. Carefully distinguishing between neurons with fragmented, intermediate, and filamentous mitochondria is critical for determining the qualitative effects of different genes in nematode excitotoxicity. This approach can also be used to discover neuron-specific vulnerability to mitochondrial perturbation.
Isolation of nematode neurons for transcriptome analysis is based on protocols developed in the Kuhn63,64, Miller57, Murphy58, and Shaham59 labs. This protocol describes sorting of all (30 neurons/animal) at-risk neurons exposed to excitotoxicity, using expression of a bright nuclear GFP in glr-1 expressing neurons54. However, single-cell -specific promoters can also be used to express the fluorescent protein markers, to obtain individual neurons of interest for cell-specific transcriptome analysis (thus circumventing problems of shallow coverage, i.e., small number of reads per gene, seen in some cases with single-cell transcriptome analysis). Sorting large quantities of neurons from excitotoxicity strains animals in the L2 stage seems more successful than when using animals in later stages. This is probably because of the increase fragility of at-risk neurons at the L3 stage, decreasing the representation of at-risk neurons following the sorting process. However, the dissociation and neuron sorting protocols can be used to sort neurons from any C. elegans larval stage.
In sum, using the nematode to illuminate key events in excitotoxicity, such as endogenous excitoprotective transcriptional programs and mitochondrial mechanisms, could potentially identify highly conserved core steps in this critical form of neurodegeneration, and might therefore eventually reveal drug targets for late stage stroke intervention.
The authors have nothing to disclose.
We thank all members of the Mano Lab and the Li lab (current and recent) for their help and support. We thank Dr. Monica Driscoll (Rutgers Univ.) for pioneering the analysis of necrotic neurodegeneration in nematodes and providing continuous support; Dr. Chris Li (CCNY) for support and advice; Jeffery Walker (CCNY Flow Cytometry Core facility), Dr. Bao Voung (CCNY), and Stanka Semova (Rockefeller Univ. Sorting Faculty Core) for practical support and advise on cell sorting; Dr. Chris Rongo (Rutgers Univ.) for reagents; Drs. David Miller (Vanderbilt Univ.), Coleen Murphy (Princeton Univ.), Shai Shaham, Menachem Katz, & Katherine Varandas (all three from Rockefeller Univ.) for C. elegans dissociation protocols.
The Mano lab received funding from NIH NINDS (NS096687, NS098350, NS116028) to I.M., and through a NIH U54 CCNY-MSKCC partnership (CA132378/CA137788).
|Cell Strainer, PluriStrainer mini 70um||PluriSelect||43-10070-40|
|Cell Strainer, PluriStrainer mini 5um||PluriSelect||43-10005-60|
|Centrifuge - 15-50 mL Sorval benchtop LEGENDX1R TC||Fisher Sci||75618382|
|Centrifuge - microfuge ; Ependorff 5424||VWR||MP022629891|
|Dry ice||United City Ice Cube|
|E. coli OP50||CGC||OP50|
|FACS tubes||USA Sci||1450-2810|
|Filter tips||USA Sci||1126-7810|
|Glass 10 mL serological pipettes||USA Sci||1071-0810|
|Immersion Oil - Carl Zeiss Immersol||Fisher Sci||12-624-66A|
|Low bind 1.5mL tubes||USA Sci||4043-1021|
|Metamorph Imaging Software||Molecular Devices|
|Microscope, Confocal, for Fluorescence Imaging||Zeiss||LSM 880|
|Microscope, Inverted, for Fluorescence Imaging||Zeiss||Axiovert 200 M|
|Microscope Camera||Q-Imaging||Retiga R1|
|Microscope Light Source for Fluorescence Imaging||Lumencor||SOLA SE Light Engine|
|Microscope, Nomarski DIC||Zeiss||Axiovert Observer A1|
|Microscope, Nomarski DIC||Nikon||Eclipse Ti-S|
|Petri dishes, 100mm||Fisher Sci||FB0875712|
|Petri dishes, 60mm||TriTech||T3308|
|Pipettor P10 Tips||USA Sci||1110-3000|
|Pipettor P1000 Tips||USA Sci||1111-2020|
|Pipettor P200 Tips||USA Sci||1110-1000|
|RNAse away spray||Fisher Sci||7000TS1|
|RNAse free serological pipettes||USA Sci||1071-0810|
|RNAse-free 50 mL tubes||USA Sci||5622-7261|
|SUPERase·in RNase inhibitor||Fisher Sci||AM2694|
|Quality Control automated electrophoresis system: Tapestation - High Sensitivity RNA ScreenTape||Agilent||5067-5579|
|Tapestation - High Sensitivity RNA ScreenTape Ladder||Agilent||5067-5581|
|Tapestation - High Sensitivity RNA ScreenTape Sample Buffer||Agilent||5067-5580|
|Tapestation - IKA MS3 vortexer||Agilent/IKA||4674100|
|Tapestation - IKA vortexer adaptor at 2000 rpm||Agilent/IKA||3428000|
|Tapestation - Loading tips||Agilent||5067- 5152 or 5067- 5153|
|Tapestation - Optical Cap 8x Strip||Agilent||401425|
|Tapestation - Optical Tube 8x Strip||Agilent||401428|
|Quality Control automated electrophoresis system: TapeStation 2200||Agilent||G2964AA|
|Tris base||Fisher Sci||BP152-500|
|Tris hydrochloride||Fisher Sci||BP153-500|
|Wescor Vapro 5520 Vapor Pressure Osmometer||Fisher Sci||NC0044806|
|Wheaton Unispense μP Dispenser||VWR||25485-003|
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