Spinal Cord Transection in the Larval Zebrafish

Summary

After spinal transection, adult zebrafish have functional recovery by six weeks post-injury. To take advantage of larval transparency and faster recovery, we present a method for transecting the larval spinal cord. After transection, we observe sensory recovery beginning at 2 days post-injury, and C-bend movement by 3 days post-injury.

Abstract

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability to reinitiate spinal neurogenesis. However, some anamniotes including the zebrafish Danio rerio exhibit both sensory and functional recovery even after complete transection of the spinal cord. The adult zebrafish is an established model organism for studying regeneration following spinal cord injury, with sensory and motor recovery by 6 weeks post-injury. To take advantage of in vivo analysis of the regenerative process available in the transparent larval zebrafish as well as genetic tools not accessible in the adult, we use the larval zebrafish to study regeneration after spinal cord transection. Here we demonstrate a method for reproducibly and verifiably transecting the larval spinal cord. After transection, our data shows sensory recovery beginning at 2 days post-injury (dpi), with the C-bend movement detectable by 3 dpi and resumption of free swimming by 5 dpi. Thus we propose the larval zebrafish as a companion tool to the adult zebrafish for the study of recovery after spinal cord injury.

Introduction

Major trauma to the human spinal cord often results in permanent paralysis and loss of sensation below the level of injury, due to the inability to regrow axons or reinitiate neurogenesis^1,2. In contrast to mammals, however, anamniotes including salamanders and zebrafish (Danio rerio) show robust recovery even after complete spinal cord transection^3,4.

The adult zebrafish is a well-established model for studying the recovery process following spinal cord injury^5-7. Following complete spinal cord transection, reestablishment of sensory and locomotive function is observed in the adult zebrafish by 6 weeks post-injury⁸. In order to examine the regenerative process in vivo, we turned to the transparent larval zebrafish⁹.

Here we present a method to transect the spinal cord of a 5 days post-fertilization (dpf) larval zebrafish using a beveled microinjection pipette as a scalpel, modified from Bhatt, et al.¹⁰ This method supports high throughput, low mortality, and reproducibility. With practice, 300 larvae/hr can be transected, and over 6 months of transections, including over 3,600 animals, 98.75% ± 0.72% survived until 7 days post-injury (dpi). Our data shows rapid recovery of sensory and locomotion as well: at 1 dpi, all movement by the injured fish is driven by pectoral fin locomotion only. However, larvae begin to respond to tungsten needle touch caudal to transection by 2 dpi, reestablish C-bend movement by 3 dpi, and display predatory swimming by 5 dpi¹¹. Using antibody staining against acetylated tubulin, we have confirmed that axons are absent from the injury site at 1 dpi, but have crossed the injury site by 5 dpi. We believe this protocol will provide a valuable technique for the study of axonal regrowth and neurogenesis in the spinal cord following injury.

Protocol

Zebrafish were raised and bred according to standard procedures; experiments were approved by the University of Utah Institutional Animal Care and Use Committee.

1. Preparation of Surgery Plates

1. Make surgery plates using 60 mm Petri dishes and Sylgard 184 Silicone Elastomer Kit, following manufacturer’s instructions. Fill dishes no more than half-full and allow to polymerize. Store covered at room temperature.

2. Preparation of Micropipettes

1. Fabricate micropipettes by heating and pulling thin-wall borosilicate capillary tubing in a micropipette puller using the same settings for making microinjection needles.
2. Under a dissection microscope, snap off tip of micropipette to approximately 200 µm in diameter with forceps.
3. Bevel broken edge with a microgrinder initially to 35°, followed by a second beveling at 25°. Ensure tip is sharp and smooth. Store finished beveled micropipette in a Petri dish on a small amount of clay.

3. Preparation of Zebrafish Larvae

1. 7 days prior to surgery, set up mating tanks of male and female zebrafish.
2. Collect embryos the following morning, 3 hr after the lights come on to ensure maximum yield. If using a transgenic reporter line such as Tg(elevl3:eGFP)^knu3, sort fertilized embryos 100/100 mm plate in 25 ml of E3 at 28.5 °C. If using wildtype, sort fertilized embryos 25/100 mm plate in 25 ml of E3 at 28.5 °C.
3. If using a reporter line, screen embryos for fluorescent expression at 48 hpf. Allow identified embryos to mature at a density of 25/100 mm plate in 25 ml of E3 at 28.5 °C.
4. When larvae are 5 dpf, prepare surgery plate by covering Sylgard with E2 + 10 mg/L Gentamycin Sulfate (GS) + Tricaine.
   1. Prepare recovery dish by adding 25 ml E2 + GS to a 100 mm Petri dish.
   2. Prepare scalpel by taping together three swabs. This will form a triangular tool with three grooves.
   3. Mount a prepared micropipette on the swabs by taping it into one of the grooves.
5. If reusing micropipettes, flush until clear with E2 + GS using a 1ml syringe and a 27 G needle prior to mounting on swabs.

4. Surgery

1. Anesthetize 1 plate of larvae at a time (25 fish) with Tricaine. Fish are sufficiently anesthetized when they no longer exhibit touch response. It is important that fish are completely anesthetized prior to surgery, otherwise they will twitch when the scalpel touches them. Surgery is performed under a dissection microscope.
2. Transfer larvae to surgery plate.
   1. Under maximum magnification, rotate one larva at a time so that it lies on its side with its back closest to the hand holding the scalpel.
   2. Position forceps so that they rest on the Sylgard, angled over the width of the larva.
   3. Bracing the glass scalpel against one of the arms of the forceps, cut into the dorsal lateral face of the larva at the level of the anal pore, being sure not to cut beyond the ventral edge of the notochord. Twist the scalpel to sever the spinal cord.
   4. Repeat with remaining larvae.
      Note: if a larva bleeds, it will not recover from the surgery. Immediately remove the larva from the surgery plate and euthanize it via Tricaine overdose.
3. Once surgery on the batch of larvae is complete, transfer injured animals to the recovery plate. This is to support the clearing of anesthesia.
   1. Caution: when collecting injured larvae for transfer, make sure they are collected head or tail first: do not stress the injury site by bending the larvae.
      Note: All devices used for surgery can be reused, including the micropipettes.

5. Recovery

1. Transfer injured larvae from the recovery plate to 100 mm plates filled with 25 ml E2 + GS at a density of 25/plate. Allow to recover in a 28.5 °C incubator.
2. Check plates daily, removing sick and dead animals. Do not change the media until Coleps (freshwater protozoa) are visible in the media. When changing the media, do not transfer the fish to a new plate; instead, remove as much media as possible and flood the same plate with new media. Repeat as necessary to reduce Coleps population.
3. Feed daily with a small amount of powdered fry food.
   Note: Live food (e.g., paramecia or rotifers) cannot be fed to injured larvae until after they have recovered locomotion. Otherwise, the live food will colonize the injury site and kill the larvae.

Results

To reduce severity of tissue damage surrounding the injury site, proper beveling of the micropipette is critical. Figure 1A shows a correctly beveled tip. Using a tip that is too wide (Figure 1B) tends to result in higher fatalities due to the increased likelihood of nicking the dorsal aorta, while a tip that is too narrow (Figure 1C) tends to glance off the skin rather than cutting tissue.

To practice this technique, it is advantageous to use...

Discussion

When initially learning this technique, we recommend attempting no more than 50-100 transections in a single session. After mastering this technique, we are able to transect up to 300 embryos per hr; however, this level of throughput requires a few months of weekly practice. We also recommend practicing with a reporter line and verifying complete transection until the incidence of incomplete spinal cord transection is reduced to less than 1%.

Spinal cord transection in the adult zebrafish...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
60 mm Petri dish	VWR	82050-544
100 mm Petri dish	VWR	89038-968
PDMS, Sylgard 184 Silicone Elastomer Kit	Fisher Scientific	NC9644388
borosilicate capillary tubing: OD 1.00 mm, ID 0.78 mm	Warner Instruments Inc.	64-0778
Forceps	Fine Scientific Tools Inc.	11252-30
Disssection microscope	Nikon	SMZ6454
Microgrinder	Narishige	EG-44
Gentamycin Sulfate	Amresco Inc.	0304-5G	dissolve in water 10 mg/ml, store at -20 °C
Tricaine	Acros Organics	118000100
Cotton tipped applicator, wood, 6-inch	Fisher Scientific	23-400-101
1 ml syringe	BD	309625
27 G needle	BD	305109
Fry food	Argent Labs	F-ARGE-PTL-CN	store at -20 °C
Micropipette puller	Sutter Instrument Co.	Model P-97	Box Filament FB330B
20x E2 (1 L); store at RT
17.5 g NaCl	Fisher Scientific	S671-500
0.75 g KCl	Fisher Scientific	P217-500
2.90 g CaCl2·2H2O	Sigma	C7902-500G
4.90 g MgSO4·7H2O	Merck	MX0070-1
0.41 g KH2PO4	Fisher Scientific	P285-500
0.12 g Na2HPO4	Sigma	S0876-500G
500x NaCO3 (10 ml); make fresh, discard extra
0.35 g NaCO3	Sigma	S5761
1x E2 (1 L); store at RT
50 ml 20x E2
2 ml fresh 500x NaCO3