Live Animal Imaging and Cell Sorting Methods for Investigating Neurodegeneration in a C. elegans Excitotoxic Necrosis Model

1. Strains used to Investigate Excitotoxic Neurodegeneration & Neuroprotection

1. 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).
2. 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.
3. Conduct genetic crosses according to standard C. elegans methods³⁰, as outlined in wormbook.org^42,43.
4. 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.
5. 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

1. Grow worms at 16-25 °C on either standard NGM plates^30,37,42 or MYOB plates^38,44 seeded with OP50 E.coli as per standard methods.
   NOTE: MYOB plates give identical results as NGM plates but are slightly easier to prepare.
2. 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

1. 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.
2. 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.
3. 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.
4. 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.
5. Identify the developmental stage of each animals by the shape of its uterus (refer to uterus diagrams in wormbook.org).
6. 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).
7. Record this data in a table where animals from a given strain are grouped by categories of developmental stage.
8. 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).
9. 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.
10. 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

1. Mount individual (or small groups of-) animals on an agar pad⁴⁶, and paralyze the worm with tetramisole (see below, section 5).
2. With a combined fluorescence and DIC scope (upright or inverted) locate specific vacuolated neurons.
3. 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

1. For imaging of mitochondrial changes in degenerating postsynaptic neurons (which are labeled with cytoplasmic GFP in the original excitotoxicity strain) examine mito-mCherry fluorescence.
2. 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.
3. 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.
4. Place animals onto the center of the tetramisole drop and mount with a coverslip.
5. Seal the sides of the coverslip with nail polish and allow the nail polish to dry.
6. Within 20 minutes of tetramisole treatment and using a scope with DIC and fluorescence imaging, locate worms using a 20x objective lens.
7. 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.
8. Capture Z-stack images using DIC, GFP, and TxRed filter settings.

6. Neuronal mitochondria morphology scoring & quantification

1. Analyze mitochondrial morphology either during live imaging of the worm or after image acquisition using ImageJ or other imaging software.
2. 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).
3. Categorize mitochondria into three main groups: filamentous, intermediate, and fragmented^50,51,52.
   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).
4. For each worm, score the percent of neurons with filamentous, intermediate, or fragmented mitochondria for a total of at least 30 worms.
5. 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

1. 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.
2. 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.
3. 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.
4. 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

1. 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.
2. Wash worms off the plates using M9 buffer. Transfer worms into 50 mL conical tubes using a glass 10 mL serological pipette.
3. Centrifuge at room temperature for 2.5 min at 250x g and remove the supernatant.
4. Resuspend the worm pellet in 10 mL of bleach solution (2% 10 N NaOH and 5% fresh household bleach in sterile deionized water).
5. 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.
   1. 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.
6. 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.
7. To retrieve the eggs/embryos, centrifuge the tubes at 250x g for 2.5 min, remove the supernatant and wash again with egg buffer.
8. Repeat four more washes.
9. 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.
10. Repeat the age synchronization by repeating this bleach/age synchronization protocol.
11. 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

1. 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.
2. 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.
3. Remove the supernatant and wash the worms with fresh M9.
4. Repeat the 30 minute gravity settling on ice.
5. Remove the supernatant, add M9 up to 45mL, and centrifuge at 250 x g for 5 minutes.
6. Transfer the pellet to a microcentrifuge tube and add M9 up to 1 mL.
7. Spin at 14,000 rpm on a tabletop centrifuge for 1 min and remove supernatant.
8. 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.
9. Add 1x egg buffer (pH 7.3, 335-345 mOsm) up to 1 mL to stop the SDS-DTT treatment.
10. Spin at 14,000 rpm on a tabletop centrifuge for 1 minute and remove supernatant.
11. To further disrupt the cuticle and dissociate the animals' cells, resuspend pellet with room temperature pronase solution, using triple the volume of the pellet.
12. Incubate at room temperature for 15-30 min.
13. 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.
14. 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.
15. In a cell culture hood, add 1x egg buffer up to 1.5 mL to stop the pronase treatment.
16. Transfer to FACS tubes, add 5mL egg buffer, and spin at 800 x g for 5 min.
17. Remove supernatant and resuspend in 3 mL pf egg buffer.
18. 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.
19. 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.
20. 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.
21. 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 lab^55,56,57, the Murphy lab⁵⁸, and the Shaham lab⁵⁹.

10. FACS machine operation modifications for identification of C. elegans neurons

1. 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.
2. 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.
3. Invert cells in Trizol 15 times to mix.
4. 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

1. For identifying GFP positive signal use the 488 nm laser with the 530/30 filter and 502 long pass (LP).
2. For identifying DAPI positive and negative cells use the 405 nm laser with the 450/50 filter.
3. 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).
4. 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.
5. Isolate foreword scatter singlets and then side scatter singlets.
6. 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.
7. Threshold for GFP+ gate is determined by comparing GFP- cells (i.e. N2) to GFP+ cells^55,56.
8. 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

1. Draw a circle on a glass microscope slide with a liquid repellent pen.
2. 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.
3. 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.

1. Thaw vials of cells in Trizol at room temperature.
2. Using a filter tip P200 pipette tip, pipette up and down several times to homogenize the sample.
3. Incubate at room temperature for 5 minutes.
4. Add 0.2 mL of chloroform per 1 mL of Trizol reagent used for lysis, then securely cap the tube.
5. Invert tubes 15 times, and incubate (at 20-25 °C) for 2-3 minutes.
6. 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.
7. 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.
8. To further purify the RNA use RNA extraction column chemistry.
9. 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.