Neuroscience
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Isolation and Culture of Oculomotor, Trochlear, and Spinal Motor Neurons from Prenatal Islmn:GFP Transgenic Mice
Chapters
Summary November 12th, 2019
This work presents a protocol to yield homogeneous cell cultures of primary oculomotor, trochlear, and spinal motor neurons. These cultures can be used for comparative analyses of the morphological, cellular, molecular, and electrophysiological characteristics of ocular and spinal motor neurons.
Transcript
This is the first protocol to simultaneously and efficiently purify and culture primary ocular and spinal motor neurons from embryonic mice. It enables the study of mechanisms underlying motor neuron disease. This protocol adds a novel in vitro oculomotor culture component to existing systems and provides pure species and age matched spinal motor neuron cultures for comparison.
In amyotrophic lateral sclerosis, spinal motor neurons degenerate while oculomotor neurons are relatively spared. Comparing these cultures could provide insights into disease mechanism and therapy. This method will facilitate studies of the mechanisms underlying motor neuron development, disease, and selective vulnerability.
Our culture system enables characterization of motor neuron morphology, molecular biology, and electrophysiology. For those new to this technique, we recommend lots of practice. Dissection can be technically challenging and multiple dissectors may be required to obtain sufficient neurons for primary culture.
Begin by harvesting Islet1 EGF positive embryos from a pregnant mouse approximately 11.5 days post-fertilization. Spray the abdomen thoroughly with ethanol and remove the uterus with sterile microdissection scissors and thumb dressing forceps. Briefly wash the uterus in sterile PBS.
Then transfer it to the dissection plate filled with pre-chilled sterile PBS. Under the bright light of the microscope, carefully remove the embryos from the uterus using microdissection scissors, thumb dressing forceps, and Dumont 5 tweezers. Then use a sterile Moria mini perforated spoon to transfer each embryo to a separate well of a 24-well plate filled with pre-chilled hibernate E low fluorescence medium supplemented with 1X V27.
While keeping the plate on ice, transfer one embryo to a sterile dissection plate and cover it completely with ice cold sterile Hank's Balanced Salt Solution or HBSS. Use tweezers to remove the face of the embryo and the tail without damaging the midbrain. Then place the embryo prone with limbs straddled and tail pointing toward the front of the microscope.
Use tweezers to slit open the roof of the fourth ventricle in order to generate a small opening. Use this opening to hook tweezers into the space created between the fourth ventricle and its roof. Dissect along the dorsal surface of the embryo rostral to the cortex and lateral to the floor plate and motor column.
Then open the dissected tissue in an open book manner to reveal the GFP positive CN3 and CN4 nuclei. Carefully separate the ventral midbrain from the embryo and remove meningeal tissue with tweezers and a microdissection knife. Dissect the bilateral GFP positive CN3 and CN4 nuclei away from the floor plate and other GFP positive surrounding tissue taking care to avoid touching or damaging the neurons.
If collecting separate CN3 and CN4 nuclei, cut along the midbrain of these two nuclei and use a P1000 pipette to collect the dissected ventral midbrain tissue with minimal HBSS. Place it in a labeled 1.7 milliliter microcentrifuge tube filled with dissection medium and store the tube on ice until dissociation. Continue pulling ventral midbrains from additional embryos in the same tube until the total number meets experimental requirements.
To dissect the ventral spinal cord, keep the embryo prone with the head facing the front of the microscope. Hold it with one pair of tweezers and insert the tip of the other pair into the unopened caudal part of the fourth ventricle. Open the rest of the hindbrain and spinal cord dorsally over the whole rostrocaudal extent of the embryo.
Cut the dorsal tissue starting from the fourth ventricle and working toward the central canal of the caudal spinal cord using the forceps as scissors. Then hold the embryo with one set of tweezers and pinch off the flap of the dorsal tissue on each side with the other pair. Remove the ventral spinal cord by using the microdissection knife to pierce directly below the GFP positive SMN lifting the ventral spinal cord with saw-like movements on both sides.
Cut the spinal cord transversely at the upper boundary of the lower limb and remove the cervical lumbar portion. Cut transversely directly above C1 where the first GFP positive anterior horn projects. Place the ventral spinal cord dorsal side up and hold it by pressing the GFP negative tissue between the GFP positive SMN columns with a pair of tweezers.
Remove the remaining attached mesenchyme, DRGs, and dorsal spinal cord by trimming both sides of the GFP positive SMN column with the microdissection knife. Use a P1000 pipette to collect the dissected ventral spinal cord tissue with minimal HBSS and place it in a labeled 1.7 milliliter microcentrifuge tube filled with dissection medium. Store it on ice until dissociation and continue pulling ventral spinal cords from additional embryos in the same tube.
Add the appropriate volume of papain solution to each of the microcentrifuge tubes with the dissected tissue samples. Incubate the tubes at 37 degrees Celsius for 30 minutes agitating the tubes every 10 minutes by finger flicking. After the incubation, gently triturate each suspension eight times with a P200 pipette and centrifuge it at 300 times g for five minutes.
Resuspend the cell pellets with the appropriate volume of albumin ovomucoid inhibitor solution by gently pipetting up and down. Repeat the centrifugation. Then carefully remove the supernatant with a P1000 pipette.
Resuspend the cells in the appropriate volume of dissection medium. Filter the suspensions through a 70 micrometer cell strainer. Next, use FACS sorting to isolate GFP positive cells dissected from CN3/CN4 and SMN.
Dilute the isolated cell suspensions with motor neuron culture medium pre-warmed to 37 degrees Celsius to the appropriate densities and add 200 microliters of the suspension to a well of a PDL laminin-coated 96-well plate. Culture the neurons in a 37 degree Celsius and 5%carbon dioxide incubator making sure to refresh the motor neuron culture medium every five days. When successfully isolated neurons were grown in culture, nearly pure primary CN3/CN4 and SMN cultures were obtained and maintained for 14 days in vitro.
The purities of the cultures at two days in vitro were 93.5%for CN3/CN4 and 86.7%for SMN when assessed by immunocytochemistry with the motor neuron marker Islet1 and neuronal marker TUJ1. The purities relied heavily on the age of the embryos and on setting the appropriate thresholds for GFP gates during FACS. The purity of neurons isolated from E10.5 embryos were comparable to those of E11.5 embryos.
However, the purity decreased significantly when E13.5 embryos were used. To determine if primary CN3/CN4s and SMNs showed differential responses to endoplasmic reticulum stressors, the cells were treated with varying concentrations of Cyclopiazonic Acid or CPA at two days in vitro and fixed three days later for immunocytochemistry to evaluate survival ratios. The number of viable neurons in each sample was counted and survival ratios were calculated.
CN3/CN4 monocultures were significantly more resistant to CPA treatment compared to SMN monocultures. Following this procedure, many additional methods can be performed to examine motor neurons. A few examples include studies of electrophysiology, cell biology, transcription, or chromatin accessibility.
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