We modified the Marmarou weight drop model for adult zebrafish to examine a breadth of pathologies following blunt-force traumatic brain injury (TBI) and the mechanisms underlying subsequent neuronal regeneration. This blunt-force TBI model is scalable, induces a mild, moderate, or severe TBI, and recapitulates injury heterogeneity observed in human TBI.
Blunt-force traumatic brain injuries (TBI) are the most common form of head trauma, which spans a range of severities and results in complex and heterogenous secondary effects. While there is no mechanism to replace or regenerate the lost neurons following a TBI in humans, zebrafish possess the ability to regenerate neurons throughout their body, including the brain. To examine the breadth of pathologies exhibited in zebrafish following a blunt-force TBI and to study the mechanisms underlying the subsequent neuronal regenerative response, we modified the commonly used rodent Marmarou weight drop for the use in adult zebrafish. Our simple blunt-force TBI model is scalable, inducing a mild, moderate, or severe TBI, and recapitulates many of the phenotypes observed following human TBI, such as contact- and post-traumatic seizures, edema, subdural and intracerebral hematomas, and cognitive impairments, each displayed in an injury severity-dependent manner. TBI sequelae, which begin to appear within minutes of the injury, subside and return to near undamaged control levels within 7 days post-injury. The regenerative process begins as early as 48 hours post-injury (hpi), with the peak cell proliferation observed by 60 hpi. Thus, our zebrafish blunt-force TBI model produces characteristic primary and secondary injury TBI pathologies similar to human TBI, which allows for investigating disease onset and progression, along with the mechanisms of neuronal regeneration that is unique to zebrafish.
Traumatic brain injuries (TBIs) are a global health crisis and a leading cause of death and disability. In the United States, approximately 2.9 million people experience a TBI each year, and between 2006-2014 mortality due to TBI or TBI sequelae increased by over 50%1. However, TBIs vary in their etiology, pathology, and clinical presentation due largely in part to the mechanism of injury (MOI), which also influences treatment strategies and predicted prognosis2. Though TBIs can result from various MOI, they are predominately the result of either a penetrating or blunt-force trauma. Penetrating traumas represent a small percentage of TBIs and generate a severe and focal injury that is localized to the immediate and surrounding impaled brain regions3. In contrast, blunt-force TBIs are more common in the general population, span a range of severities (mild, moderate, and severe), and produce a diffuse, heterogeneous, and global injury affecting multiple brain regions1,4,5.
Zebrafish (Danio rerio) have been utilized to examine a wide range of neurological insults spanning the central nervous system (CNS)6,7,8,9. Zebrafish also possess, unlike mammals, an innate and robust regenerative response to repair CNS damage10. Current zebrafish trauma models use various injury methods, including penetration, excision, chemical insult, or pressure waves11,12,13,14,15,16. However, each of these methods utilizes an MOI that is rarely experienced by the human population, is not scalable across a range of injury severities, and does not address the heterogeneity or severity-dependent TBI sequela reported after blunt-force TBI. These factors limit the use of the zebrafish model to understand the underlying mechanisms of the pathologies associated with the most common form of TBI in the human population (mild blunt-force injuries).
We aimed to develop a rapid and scalable blunt-force TBI zebrafish model that provides avenues to investigate injury pathology, progression of TBI sequela, and the innate regenerative response. We modified the commonly used rodent Marmarou17 weight drop and applied it to adult zebrafish. This model yields a reproducible range of severities ranging from mild, moderate, to severe. This model also recapitulates multiple facets of human TBI pathology, in a severity-dependent manner, including seizures, edema, subdural and intracerebral hematomas, neuronal cell death, and cognitive deficits, such as learning and memory impairment. Days following injury, pathologies and deficits dissipate, returning to levels resembling undamaged controls. Additionally, this zebrafish model displays a robust proliferation and neuronal regeneration response across the neuroaxis concerning injury severity.
Here, we provide details toward the set up and induction of blunt-force trauma, scoring post-traumatic seizures, assessment of vascular injuries, instructions on preparing brain sections, approaches to quantifying edema, and insight into the proliferative response following injury.
Zebrafish were raised and maintained in the Notre Dame Zebrafish facility in the Freimann Life Sciences Center. The methods described in this manuscript were approved by the University of Notre Dame Animal Care and Use Committee.
1. Traumatic brain injury paradigm
- Add 1 mL of 2-phenoxyethanol to 1 L of system water (60 mg of Instant Ocean in 1 L of deionized RO water).
- Prepare an aerated recovery tank containing 2 L of system water at room temperature.
- Select the desired weight of ball bearing and a desired length and diameter of steel/plastic tubing and determine the energy and impact force.
NOTE: Tubing should have an inner diameter that allows the ball bearing to pass through without changing its path or speed of movement.
- Determine the kinetic energy upon impact:
Where, KE = kinetic energy, m = mass (in kg), g = gravitational force, h = height (in m) from drop point to fish.
NOTE: This provides kinetic energy in J. Multiply the value by 1,000 to determine mJ. KE is based on an accelerating object, which occurs when the ball bearing is dropped from a stationary position.
- Generate a mild TBI (miTBI) using 7.62 cm long steel/plastic tubing that ends 1.5 cm above the plate on the zebrafish skull (total distance 9.1 cm) and a 1.5 g (6.4 mm diameter) ball bearing. These produce a kinetic energy of 1.33 mJ. This damage was empirically decided to be equivalent to a miTBI based on key pathophysiological TBI markers, such as vascular injury, subdural/intracerebral hematoma formation, neuronal cell death, and cognitive impairments that largely recapitulated what was reported in the human population in a severity-dependent manner.
- Calculate kinetic energy miTBI =
- Generate a moderate TBI (moTBI) using 12.7 cm long steel/plastic tubing that ends 1.5 cm above the plate on the zebrafish skull (total distance 14.2 cm) and a 1.5 g (6.4 mm diameter) ball bearing that produces kinetic energy of 2.08 mJ.
- Calculate kinetic energy moTBI =
- Generate a severe TBI (sTBI) using 7.62 cm long steel/plastic tubing that ends 1.5 cm above the plate on the zebrafish skull (total distance 9.1 cm) and a 3.3 g (8.38 mm diameter) ball bearing that produces a kinetic energy of 2.94 mJ.
- Calculate kinetic energy sTBI =
- Determine the kinetic energy upon impact:
- Fill a Petri dish with modeling clay (Figure 1, step 1) and use a blunt instrument (i.e., the back of a pair of forceps) to create a raised platform (5 cm x 1.5 cm) with additional modeling clay (Figure 1, step 2).
- Use a razor blade to divide the raised platform lengthwise into two approximately equal halves (Figure 1, step 3, red dashed line). Form the two halves into a channel that accommodates the length of an adult fish (Figure 1, step 4). Use additional clay to build walls that will secure ~2/3 of the fish body, leaving the head exposed.
- Mold a small support in the exposed head region perpendicular to the walls to support the head to avoid rotation or recoil of the head upon injury (Figure 1, step 4).
NOTE: The channel should be deep enough to support the fish in a dorsal-up position but should still allow the head to rest above the surrounding clay. Additionally, the head support should follow the natural curvature of the fish, supporting the lower mandible and gills.
- Ensure that the dropped weight is not impeded by the channel's sides (Figure 1, step 4).
- Create a 3 mm diameter steel disk using a mini hole-punch and 22 g steel flashing. Each disk may be used multiple times.
- Anesthetize one fish by placing it into a beaker with 50-100 mL of 1:1000 (1 mL/1 L, 0.1%) 2-phenoxyethanol until it is unresponsive to tail pinch.
- Place the fish, dorsal side up, on the clay mold within the channel so that the body is secured on the sides and place a 3 mm 22 g steel disc on the head, centered over the desired impact point (Figure 1, step 5).
- Ensure that the fish is aligned as perpendicularly as possible to avoid its head from tilting to one side, which could cause an uneven impact.
- Secure the steel/plastic tubing, using a standard ring stand and arm clamp, so the bottom of the tubing is 1.5 cm above the head of the zebrafish (Figure 1, step 6). Ensure the tubing is straight.
- Look down the tubing and ensure that the tubing is aligned above the steel plate.
- Drop the ball bearing (1.5 g for mild and moderate TBI, and 3.3 g for severe TBI) from a predetermined height (described in Steps 1.3.3-1.3.5) down the tubing onto the steel plate located over the desired neuroanatomical region of interest (e.g., cerebellum, Figure 1, step 5) to produce the blunt-force TBI for the desired severity of injury. Place the injured fish in a recovery tank to be monitored.
NOTE: Depending on the severity of the injury, mortality or tonic-clonic seizures may occur. If the fish are unresponsive for a prolonged period of time following the TBI, use a transfer pipette or forceps to administer a tail-pinch and evaluate for a pain response.
2. Scoring seizures post-TBI in the adult zebrafish
- Anesthetize and injure the fish according to the injury protocol outlined in section 1 and place the injured fish in an aerated recovery tank of 2 L of system water.
- Observe the fish for any signs of post-traumatic seizures beginning immediately after being placed in the recovery tank. Set an observation time (i.e., 1 h) and record all seizure activity, including the seizure score (described below), duration of each seizure, and the percentage of fish that experienced seizures (Figure 2A).
NOTE: Seizures can occur immediately upon injury, as well as hours or days post-TBI. Seizures can persist long term and a single fish can have multiple seizures. Setting an observation time of 1 h or greater will yield a good representation of the overall trend of seizure rates.
- Score fish using the Mussulini18 guidelines for adult zebrafish seizure phenotypes.
NOTE: Without using the tracking software, it is difficult to assess seizure scores below 3 unbiasedly. Therefore, only seizure scores of 3 or greater should be recorded when the assessment is being performed without software.
3. Brain dissection
- Euthanize fish in a 1:500 (2 mL/1 L, 0.2%) solution of 2-phenoxyethanol, until gill movements cease, and they are unreactive to fin pinch, at the desired endpoint.
- Fill a Petri dish with modeling clay and create a small cavity to support the body during dissection.
- Place the fish, dorsal side up, in the clay mold. Place one dissection pin through the midline halfway down the body of the fish and a second pin ~5 mm behind the base of the head.
- Under a dissecting light microscope, bluntly sever the optic nerve with a pair of #5 Dumont forceps and remove the eyes (Figure 2A).
- Orientate the fish so that the rostral end is furthest when looking through the microscope (Figure 2B).
NOTE: The following steps are for right-handed individuals. Left-handed individuals may prefer to perform the following steps in a mirrored orientation.
- Use #5 forceps to slowly place one end of the forceps under the right parietal plate, making a deliberate scissor action moving toward the rostral end and remove the right parietal and frontal plates (Figure 2C).
- Keep the forceps at an angle of 45° or lower in order to avoid penetrating the brain during dissection.
- Rotate the fish 90˚ clockwise. Place one end of the #5 forceps under the left parietal plate and use the same scissor motion to remove the left parietal and frontal plates exposing the entire dorsal aspect of the brain (Figure 2D,E).
- Bluntly transect the maxilla with #5 forceps. Preserve the olfactory bulbs and do not damage them if this is the region of interest.
- Remove the right opercle, preopercle, interopercle, and subopercle with #5 forceps (Figure 2F).
- Bluntly resect the musculature at the caudal end of the calvarium opening using #5 forceps, exposing the spinal cord.
- Bluntly transect the spinal cord with #5 forceps. Carefully place forceps under the brain and gently remove the brain from the calvarium.
NOTE: Never pinch the brain. Use forceps to "cradle" the brain or resect caudally and utilize the exposed spinal cord as a pinch point to maneuver the brain.
- Fix the removed brains in 9 parts 100% ethanol to 1 part 37% formaldehyde overnight at 4°C on a rocker platform.
4. Edema studies in the zebrafish brain
- Anesthetize and injure fish according to the injury protocol outlined in section 1 and allow the fish to recover in a recovery tank until they start swimming freely.
- Place fish back in the normal housing conditions post-injury for 1 day.
- Euthanize the fish in 1:500 2-phenoxyethanol, after time has elapsed.
- Dissect the whole brain or region of interest according to the protocol outlined in section 3 and place the brain immediately on a small weigh boat.
NOTE: Use caution when transferring the brain, using fine forceps to gently place it on the weigh boat without stabbing or scraping the brain, which could result in the loss of tissue.
- Label (with injury group and brain number) and tare an additional small drying weigh boat on a scale. Use a scale with the ability to measure a minimum of 0.001 g to get an accurate measurement.
- Transfer the brain to the tared drying weigh boat and record the wet weight of the brain. Orient the brains so that they lay flat on the weigh boat with the dorsal side facing up.
- Place the drying weigh boat and brain in a hybridization oven set to 60 °C for 8 h.
- After drying, the brain may stick to the drying weigh boat and may be difficult to remove and transfer to a new tared small weigh boat. Avoid pinching the brain with forceps, as this might result in damage to the dry brain and loss of tissue. Instead, pinch the fine forceps together and, starting at the ventral side of the brain, scoop in an upward motion.
- Determine the water content of each brain using the formula (Figure 4):
5. Labeling cellular proliferation across the neuroaxis and preparing fixed tissue.
- Prepare 10 mM 5-ethynyl-2'-deoxyuridine (EdU) in 2 mL of ddH2O.
- Anesthetize 3 to 4 fish at a time in 50-100 mL of 1:1000 (1 mL/1 L, 0.1%) 2-phenoxyethanol until the fish are unresponsive to tail pinch, at the desired time point following injury (using the protocol outlined in section 1).
- Make a partial incision on a wet sponge and place one fish at a time in the opening, ventral side up.
- Use a 30 G needle to inject ~40 µL of 10 mM EdU into the fish's body. Return the fish to a holding tank filled with system water.
NOTE: Repeated injections may be performed at different time points to label a greater number of proliferating cells and may be needed if a chase period of greater than 1 week is desired.
- Collect brains as outlined in section 3 and place them as a group in a 5 mL glass vial containing 2 mL of 9 parts 100% ethanol to 1 part 37% formaldehyde. Fix the brains at 4 °C on a rocker platform.
NOTE: A rocker or shaker platform prohibits the brains from resting at the bottom and the telencephalon of whole brains from curling.
- Rehydrate brains, in the same glass vial used to fix brains, in washes of descending ethanol series, 75%, 50%, and 25%, for 15 mins each, followed by a 1.5 h wash in 5% sucrose/PBS on a rocker at room temperature. Store the brains in a glass vial overnight in 30% sucrose/PBS at 4° C on a rocker platform.
- Remove brains from 30% sucrose/PBS and transfer them with forceps into a 12-well plate (one treatment group per well) with wells filled with a 2:1 solution consisting of 2 parts tissue freezing medium and 1 part 30% sucrose/PBS. Incubate the brains overnight at 4 °C on a rocker platform.
- Transfer brains to the next row of wells within the 12-well plate submerging brains in 100% TFM for 2-24 h at 4 °C.
- Use a cryostat chuck to embed the brains in TFM in the desired orientation on dry ice.
- Perform cryosectioning (16 μm thick sections) and collect the sections on positively charged slides. Dry slides on a slide warmer for 1 h and then store at -80 °C or continue with immunohistochemistry.
- Prepare a hydrophobic barrier on the slide around the tissue sections and allow to dry on a slide warmer for 20 min.
- Briefly wash the slides in PBS for 5 min and then twice in PBS-Tween 20 (0.05%) for 10 min each.
- Perform EdU detection using the EdU Cell Proliferation Kit and the manufacturer's instructions.
- Analyze the slides and quantify the fluorescent EdU-labeled cells using either an epifluorescent microscope or a confocal microscope. A minimum of a 40x objective will be required to clearly distinguish individual cells.
Preparing the injury-induction rig allows for a rapid and simplistic means of delivering a scalable blunt-force TBI to adult zebrafish. The graded severity of the injury model provides several easily identifiable metrics of successful injury, though the vascular injury is one of the easiest and most prominent pathologies (Figure 3). The strain of fish used during the injury can make this indicator easier or harder to identify. When using wild-type AB fish(WTAB, Figure 3A-D), identification of vascular injury can be difficult to distinguish between either miTBI or moTBI and undamaged control fish due to the pigmentation (Figure 3A-C). Following injury, miTBI fish display minimal surface abrasions (Figure 3B), while moTBI exhibit limited cerebral hemorrhaging (Figure 3C). While sTBI can still be challenging, the extent of injury is often apparent (Figure 3D). In contrast, when using albino (Figure 3E-H) or casper fish (Figure 3I-L), vascular injury is easily identified. Additionally, impact seizures are often observed following injury and seizure rate among the group is another representative metric of injury (Table 1). Injured fish will display tonic-clonic seizures (ataxia, ZBC 1.9, bending, ZBC 1.16, circling, ZBC 1.32, and corkscrew swimming, ZBC 1.37)19 that are easily observed following injury, regardless of the background strain. Seizures will be observed with increasing prevalence in relation to severity. After injury, miTBI do not display seizure-like behaviors; however, moTBI will display seizure behaviors (10.66% ± 1.37%, p < 0.0001, Table 1) and the incidence is further elevated in sTBI fish (19.93% ± 1.49%, p < 0.0001, Table 1).
Successful removal of the brain is critical for a myriad of further investigations, such as edema and assessing cell proliferation. Perform dissections with the utmost care to avoid damaging brain regions (most often by unintentional puncture) and to conserve all regions (the olfactory bulbs can easily be lost). Following the brain dissection procedure and schematic (section 3, Figure 2A-F) allows for a complete brain removal (Figure 2G,H). Investigators should consider whether their analysis requires the entire brain or whether a collection of specific brain regions may suit their needs. Dependent on the severity of the injury and time of collection, the brains may exhibit attached subdural hemorrhages, however, these are often adhered to the underside of the skull and lost during dissection. Swelling of the cerebrum is at times apparent, but due to anatomical differences and variation in general size, edema is the best method to assess swelling. Following the protocol outlined (section 4), undamaged brains exhibit a fluid content of 73.11% ± 0.80%, and miTBI, though slightly elevated, do not display a significant increase in edema at 1, 3, or 5 dpi (1 dpi: 76.33% ± 1.32%, p = 0.36, 3 dpi: 75.33 ± 1.37%, p = 0.84, 5 dpi: 74.14 ± 1.50%, p > 0.99, Figure 4). In contrast, both moTBI and sTBI had significant edema 1 dpi (moTBI: 80.55 ± 0.94%, p < 0.0001, sTBI: 86% ± 1.05%, p < 0.0001), and 3 dpi (moTBI: 78.11 ± 0.93%, p < 0.018, sTBI: 77.77% ± 1.02%, p < 0.036, Figure 4). However, fluid content of both moTBI and sTBI returned to levels resembling undamaged controls by 5 dpi (moTBI: 74.42 ± 1.25%, p > 0.99, sTBI: 73.85% ± 1.01%, p > 0.99, Figure 4).
Cell proliferation, following TBI in zebrafish, is a robust assessment of the extent of injury. While the cell proliferation response has been studied previously in zebrafish following other forms of brain injury9,12, in most instances, the investigation was limited to the injury site. This blunt-force TBI results in a robust proliferation response spanning the neuroaxis. In a severity-dependent manner (sTBI data shown), increased EdU labeling is observed in the ventricular and subventricular zones of the forebrain (telencephalon, Figure 5B) relative to undamaged controls (Figure 5A). As sections moved caudally into the midbrain (mesencephalon and diencephalon), injured brains displayed increased EdU labeling in the periventricular grey zone (PGZ) the optic tectal lobes (TeO), and aspects of the anterior hypothalamus compared to undamaged fish (Figure 5D and Figure 5C, respectively). In the hindbrain, neurogenic regions that are evident in the undamaged brain (Figure 5E,G) exhibit increased cell proliferation following the sTBI (Figure 5F,H).
To summarize, a modified Marmarou weight drop applied to adult zebrafish provides a reproducible and scalable mild, moderate, or severe blunt-force TBI. Zebrafish, in a severity-dependent manner, display various pathologies, including seizures and vascular injury (i.e., subdural and intracerebral hematomas). Additionally, injured fish display decreased recovery rate (analogous to loss of consciousness, cognitive deficits in the form of learning and memory issues, and neuronal cell death (data not shown). The pathologies observed, rapidly recover over the span of 4-7 days coinciding with robust proliferative events across the neuroaxis.
Figure 1: Setting up the scalable injury apparatus. Graphical representation of the setup, the model, and delivery of scalable TBIs to the zebrafish. Steps 1-4 provide instructional overview of the steps to form the support mold that immobilizes the fish and exposing the head during damage. Steps 5-7 provide instructions on delivering the injury with insight on aspects to consider when troubleshooting the model. The figure was created with BioRender.com. Please click here to view a larger version of this figure.
Figure 2: Skull removal for brain dissection. Schematic of a simplified zebrafish skull and the step-by-step removal of bone (blue sections) to expose the adult zebrafish brain. (A,B) The eyes are bluntly removed with #5 forceps severing the optic nerves. (C) Forceps are placed into the musculature directly caudal to the parietal plates (black arrow) to remove the right parietal bone and then the right frontal bone. (D,E) The left parietal bone and left frontal bone are removed. (F) The right opercle, preopercle, interopercle, and subopercle are removed, providing lateral and dorsal access to the brain. (G,H) Undamaged and sTBI brains were removed. Scale bar = 500 µm. Please click here to view a larger version of this figure.
Figure 3: Vascular injury in various backgrounds across injury-severities. Dorsal view of undamaged and TBI wild-type AB, albinob4, and casper adult zebrafish displaying vascular injury. (A-D) Adult wild-type AB fish are heavily pigmented and abrasions following miTBI (B) are difficult to visualize. Vascular injury was more apparent in moTBI (C) and sTBI (D) fish compared to undamaged controls (A). (E-H) albino fish were less pigmented and visualization of the brain was more distinct. Vascular injury following TBI was clearly observed and distinguished across severities. (I-L) casper fish provided the most adaptable background for novice investigators as the transparency allowed for easy identification of desired neuroanatomical regions and clear observation and delineation of vascular injury by TBI-severity. Scale bar = 500 µm. Please click here to view a larger version of this figure.
Figure 4: Zebrafish experience injury-induced edema following TBI. Zebrafish were exposed to different severities of TBI (undamaged, miTBI, moTBI, and sTBI) and assessed at different days following damage for percent fluid content (edema). Statistical analyses were performed with a Browns-Forsythe and Welch ANOVA followed by a Dunnett's T3 multiple comparison post-hoc test. n = total number of individual fish. All statistical analyses were performed with the Prism (Graphpad 9.0) software package. Please click here to view a larger version of this figure.
Figure 5: TBI-induced proliferation across the neuroaxis. (A-H) Confocal images of coronal and sagittal brain sections of undamaged and sTBI fish that were IP-injected with EdU 12 h prior to collection. Increased EdU incorporation was observed in multiple neurogenic niches following injury across the forebrain (A,B), midbrain (C,D), and hindbrain (E-H). Cerebellum, CCe, Granule Layer, GL, Medial Valvula Cerebelli, Vam, Molecular Layer, ML, Optic Tectum, TeO, Periventricular Grey Zone, PGZ, Telencephalon, and Tel. All scale bars are 200 µm. Please click here to view a larger version of this figure.
|Group||N||n||Mean Seizures (%) ± SEM||p|
|moTBI||10||184||10.66% ± 1.37%||<0.0001|
|sTBI||10||237||19.93% ± 1.49%||<0.0001|
Table 1. Zebrafish display severity-dependent impact seizures following TBI. Quantification of tonic-clonic seizures, which was recorded as the percent of an experimental injured group, that were observed within 1 h post-injury. Statistical analyses were performed with a one-way ANOVA followed by Tukey's post-hoc test. N = total number of experimental groups, n = total number of individual fish. Statistical analyses were performed using the Prism (Graphpad 9.0) software package.
Investigations of neurotrauma and associated sequelae have long been centered on traditional non-regenerative rodent models20. Only recently have studies applied various forms of CNS damage to regenerative models9,11,13,14,21. Though insightful, these models are limited by either their use of an injury method uncommonly seen in the human population (penetrating traumas, chemical ablation, blast) and/or the injury is not scalable and therefore does not fully address the heterogeneity of severity-dependent pathologies observed in the human population22,23,24,25. Here, we provide a damage paradigm that applies the most common form of clinically relevant head trauma (blunt-force)4 that produces many of the pathological metrics established in human diagnosis22,23,24,25. When applied to the regenerative zebrafish, the model provides avenues to investigate the progression and recovery of injury-induced pathologies across severities, such as edema or post-traumatic seizures, as well as elucidating the mechanisms behind the innate regenerative recovery.
There are two key features of our model in producing a blunt-force TBI in zebrafish. First, our model delivers an inexpensive and simple injury paradigm that is rapid, which permits the successive injury to a large number of individuals or repeated injury to an individual to investigate the cumulative effect of blunt-force TBI. Second, this model is easily scalable to examine the effects of different force impacts. By changing the length of the tube (the height from which the ball bearing is dropped) and the weight of the ball bearing, the energy delivered to the skull of the fish, and the impact force can be easily modified and computed. This scalability of the injury allows for multiple avenues of investigation in regard to severity-dependent TBI sequelae progression and regenerative mechanisms of CNS repair.
With multiple metrics to access successful injury application, careful consideration should still be given to the genetic background of fish to be utilized. The casper or albino mutant fish will be favorable for novice investigators to reliably place the fish under the drop shaft, the placement of the steel disk over the desired neuroanatomical impact point, and assessing vascular injury. Furthermore, careful removal of the brain is simplified by the visual accessibility of the bones and brain in the casper and albino mutant fish. However, pigmented wild-type fish can be used though identification of landmarks and successful dissection may come with notable practice. Furthermore, pigmented fish can be used when producing either a moTBI or sTBI insult, as the subsequent pathologies allow for proper characterization of injury.
One major reason to study the effects of blunt-force TBI in zebrafish is to examine the source of injury-induced cell proliferation and the mechanisms underlying neuronal regeneration. Developmental and basal levels of constitutive proliferation have been identified in neurogenic niches across the zebrafish neuroaxis26,27, and injury-induced regeneration has been observed localized or adjacent to the injury site in adult zebrafish8,12,15. However, our blunt-force TBI model demonstrates that the diffuse injury also results in a severity-dependent cell proliferation event within the neurogenic niches across the neuroaxis. Identification of the source and extent of cell proliferation following TBI will permit the application of single-cell RNA-Seq to identify the changes in gene expression in the proliferative niche and testing the role of different signaling pathways, through the application of specific agonists and antagonists, in regulating this regeneration response. This approach has proven useful in elucidating the mechanisms underlying neuronal regeneration in the damaged zebrafish retina28 and should be equally useful in the brain following TBI.
In conclusion, our model provides a rapid, simple, and cost-effective injury method to deliver a scalable blunt-force TBI. This model will be useful to further investigate the effects of severity-dependent or repeated blunt-force TBI, as well as elucidating therapeutic targets of genetic regulation improving neuronal protection or inducing neuronal regeneration for functional cognitive recovery in adult vertebrates.
The authors have nothing to disclose.
The authors would like to thank the Hyde lab members for their thoughtful discussions, the Freimann Life Sciences Center technicians for zebrafish care and husbandry, and the University of Notre Dame Optical Microscopy Core/NDIIF for the use of instruments and their services. This work was supported by the Center for Zebrafish Research at the University of Notre Dame, the Center for Stem Cells and Regenerative Medicine at the University of Notre Dame, and grants from National Eye Institute of NIH R01-EY018417 (DRH), the National Science Foundation Graduate Research Fellowship Program (JTH), LTC Neil Hyland Fellowship of Notre Dame (JTH), Sentinels of Freedom Fellowship (JTH), and the Pat Tillman Scholarship (JTH).
|#00 buckshot||Remington||RMS23770||3.3g weight for sTBI|
|#3 buckshot||Remington||RMS23776||1.5g weight for miTBI/moTBI|
|#5 Dumont forceps||WPI||14098|
|5ml glass vial||VWR||66011-063|
|Click-iT EdU Cell Proliferation Kit||Life Technologies||C10340|
|CytoOne 12-well plate||USA Scientific||CC7682-7512|
|Instant Ocean||Instant Ocean||SS15-10|
|Super frost postiviely charged slides||VWR||48311-703|
|Super PAP Pen Liquid Blocker||Ted Pella||22309|
|Tissue freezing medium||VWR||15148-031|
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