A Repetitive Weight-Drop Procedure for Establishing a Mouse Model of Chronic Traumatic Brain Injury

Summary

A protocol is presented for establishing a repetitive closed-head mild traumatic brain injury mouse model for studying the mechanisms underlying chronic depressive-like behavior in the absence of gross structural brain lesions.

Abstract

Traumatic brain injury (TBI) can lead to persistent neuropathological changes and long-term neuropsychiatric sequelae. During recovery, a substantial proportion of patients develop affective disturbances, including depressive symptoms and sleep-related complaints, even when routine computed tomography or magnetic resonance imaging reveals no obvious structural abnormalities. Experimental models that reproduce these features are needed to investigate the mechanisms underlying chronic post-traumatic neuropsychiatric symptoms. Here, we present a protocol to establish a repetitive closed-head mild traumatic brain injury mouse model for studying the mechanisms underlying chronic depressive-like behavior in the absence of overt gross structural brain lesions detectable by macroscopic examination and T2-weighted MRI. In this method, the intact skull is exposed and subjected to repeated impacts using defined weight-drop parameters over five consecutive days. Under the conditions described here, the procedure produces a closed-head injury without gross skull fracture or macroscopic focal contusion. After a post-injury interval, mice exhibit depressive-like behaviors while showing no obvious focal structural abnormalities on gross examination or T2-weighted magnetic resonance imaging under the conditions used in this study. This protocol provides an experimental approach for studying chronic depressive-like behavior after repetitive mild traumatic brain injury and for evaluating potential therapeutic interventions.

Introduction

However, because the present paradigm primarily delivers linear impact to a head-fixed mouse, it should be interpreted as a closed-head rmTBI model optimized for studying chronic affective outcomes rather than as a direct biomechanical mimic of sports-related concussion or minor traffic injury.”Traumatic brain injury (TBI) is currently one of the leading causes of trauma-related morbidity and mortality, imposing a substantial medical and economic burden on families and society^1,2. Notably, central nervous system damage following TBI is not transient; it typically evolves in a chronic and progressive manner. A study on TBI revealed that even 18 years post-injury, microglial activation persists, and prolonged chronic neuroinflammation continues to cause severe neural deterioration³. Epidemiological data indicate that up to 28%–32% of patients develop severe affective disorders during the prolonged recovery phase⁴. In this article, rmTBI refers to the repetitive mild traumatic brain injury induction paradigm, whereas the subsequent chronic post-injury stage, characterized by persistent behavioral alterations, is referred to here as chronic traumatic brain injury (c-TBI). The term c-TBI is used here to describe the chronic post-injury state characterized by persistent behavioral alterations after rmTBI, rather than a distinct injury paradigm. This usage is conceptually consistent with our previous work demonstrating that experimental TBI can evolve as a progressive chronic process associated with long-term histopathological and behavioral abnormalities.

These chronic neuropsychiatric sequelae severely compromise patients’ quality of life and functional outcomes⁵. However, the exact pathogenesis underlying these secondary affective disorders remains elusive, and clinical treatment options are severely limited. Therefore, investigating the mechanisms of c-TBI-induced depressive-like behaviors and identifying precise therapeutic targets is of urgent clinical need and significant practical importance.

To elucidate the pathological mechanisms of TBI, researchers have developed various animal models, among which the controlled cortical impact (CCI) model is currently one of the most widely used^6,7. CCI allows for the precise control of impact parameters to generate highly reproducible focal lesions. However, both CCI and traditional fluid-percussion injury (FPI) models typically require a craniotomy. These approaches are particularly useful for studying focal contusion, tissue loss, and severe lesion-related neuroinflammatory responses, but they tend to simulate severe open TBI, which significantly differs from the most common clinical presentations of mild closed-head injuries, such as sports-related concussions^8,9. Crucially, the brain tissue necrosis, glial scarring, and late-stage cavitation caused by craniotomy and impact present two major limitations. First, severe structural deficits often induce motor dysfunction, which directly confounds the accurate assessment of behavioral outcomes. Second, the physical loss of tissue severs long-range neural projections. As demonstrated in previous studies using the CCI-induced TBI model, mice exhibited significant focal brain contusions⁴. This structural disruption severely impedes the application of advanced neuroanatomical techniques, such as viral tracing, making it largely unfeasible to investigate psychiatric symptoms and their associated neural circuits. Accordingly, although these models remain valuable for studies of severe or focal TBI, they may be less suitable when the goal is to investigate chronic affective abnormalities after repetitive mild closed-head injury, especially when preservation of structural integrity is important for behavioral and circuit-level analyses.

To overcome the limitations of the aforementioned severe open TBI models, this study details a protocol for establishing a repetitive mild traumatic brain injury (rmTBI) murine model using a weight-drop method on the exposed, intact skull without craniotomy. This protocol employs precisely defined impact parameters: a drop height of 25 cm and a weight of 20 g. The impact site was targeted at 2 mm left of the sagittal suture and 2 mm posterior to the coronal suture, using a curved metal impactor tip with a diameter of 4 mm to ensure the accuracy and uniformity of the impact force. Furthermore, the specific frequency of repetitive impacts over five consecutive days ensures the stability of chronic pathological alterations. Comprehensive evaluations incorporating MRI, histopathology, and behavioral assays demonstrate that this murine model exhibits significant depressive-like behaviors while preserving gross structural brain integrity, without obvious focal contusions or motor deficits. This phenotypic profile captures selected behavioral features of repetitive mild head injury, including neuropsychiatric-like changes despite no obvious structural abnormalities on routine neuroimaging^10,11,12,13. However, because the present paradigm primarily delivers linear impact to a head-fixed mouse, it should be interpreted as a closed-head rmTBI model optimized for studying chronic affective outcomes rather than as a direct biomechanical mimic of sports-related concussion or minor traffic injury.

Protocol

All animal procedures were approved by the Institutional Animal Care and Use Committee of the Institute of Artificial Intelligence, Hefei Comprehensive National Science Center (Approval No. IAI2024071004). All procedures were performed in accordance with institutional and national guidelines for the care and use of laboratory animals. A schematic diagram of the experimental protocol is shown in Figure 1. Equivalent equipment from other vendors may be used, provided that the critical performance characteristics are maintained, such as the 20 g impact weight, 25 cm drop height, 4 mm impactor tip diameter, reliable isoflurane delivery, and MRI settings comparable to those used here. The reagents and the equipment used are listed in the Table of Materials.

1. Animal preparation

1. Prepare the animals.
   1. Use male C57BL/6N mice aged 8–10 weeks and weighing 20–25 g.
   2. House the mice in individually ventilated cages (IVCs) in a specific pathogen-free (SPF) animal facility. Provide food and water ad libitum and maintain a 12 h light/12 h dark cycle under controlled temperature and humidity.
   3. Acclimate the mice to the housing room before the experiment and perform routine health screening in accordance with institutional animal care procedures.
2. Assign the groups.
   1. Randomly assign the mice to the Naive group, the Sham group, and the c-TBI group. Investigators responsible for MRI interpretation, gross pathological assessment, and behavioral scoring/data analysis were blinded to group assignment.
   2. Record the animal identification number and group assignment before the first injury session.

2. Preparation for surgery and anesthesia

1. Prepare the surgical instruments and anesthesia system.
   1. Prepare a sterile scalpel fitted with a No. 11 blade, sterile cotton swabs, sterile physiological saline, 10% povidone-iodine solution, 5-0 nonabsorbable polypropylene monofilament suture, adhesive tape, and chlortetracycline ophthalmic ointment.
   2. Sterilize the surgical instruments before use, place them on a clean surgical field, and maintain aseptic technique throughout the procedure. Wear appropriate personal protective equipment, including gloves, a lab coat, and eye protection.
   3. Prepare a thermostatic heating pad and set it to 37 °C ± 0.5 °C.
   4. Connect the induction chamber and nose cone to the laboratory animal gas anesthesia system.
   5. Set the oxygen flow rate to 0.5– 1.0 L/min and prepare the isoflurane vaporizer for induction and maintenance. Perform anesthesia in a well-ventilated area or use an approved waste-gas scavenging system.
2. Prepare the mouse for surgery.
   1. Administer buprenorphine HCl (0.1 mg/kg, intraperitoneally) 30 min before anesthesia and surgery in accordance with the approved animal protocol.
   2. Place the mouse in the induction chamber and induce anesthesia with isoflurane. Set the vaporizer to 3%–5% for induction and monitor the mouse continuously until the righting reflex is lost and spontaneous limb movement ceases. Do not exceed 5% isoflurane during induction.
   3. Transfer the mouse to a stereotaxic frame in the prone position and maintain anesthesia through a nose cone. Reduce the vaporizer setting to 1%–2% for maintenance while keeping the oxygen flow rate at 0.5–1.0 L/min.
      1. During the procedure, maintain a stable respiratory rate of approximately 60–100 breaths/min. If the respiratory rate falls below 60 breaths/min or breathing becomes shallow or irregular, immediately reduce the isoflurane concentration, provide supportive warming, and delay impact until stable respiration resumes. Do not exceed 2% isoflurane during maintenance.
   4. Confirm adequate anesthesia by the absence of a pedal withdrawal reflex, secure the head with the ear bars of the stereotaxic frame without damaging the tympanic membranes (Figure 2A), and maintain the mouse in the prone position. If adequate anesthesia cannot be achieved within the stated induction and maintenance ranges, or if excessive respiratory depression occurs, exclude the animal from that injury session.
   5. Apply chlortetracycline ophthalmic ointment to both eyes, shave the scalp, and disinfect the surgical site with 10% povidone-iodine solution.

3. Establishing the repetitive mild traumatic brain injury model

1. Expose the skull.
   1. Make a longitudinal midline incision, approximately 0.5 cm to 1 cm in length, in the scalp using a sterile scalpel fitted with a No. 11 blade.
   2. Bluntly dissect the subcutaneous tissue and periosteum with sterile forceps or cotton swabs to fully expose the skull without scratching the bone surface.
   3. Wipe the skull surface with a sterile cotton swab moistened with physiological saline to visualize the cranial landmarks, including bregma and the coronal suture.
2. Prepare the impact device.
   1. Install a 4 mm diameter curved metal impactor tip on the free-falling weight-drop device.
   2. Set the drop height to 25 cm and use a 20 g impact weight (Figure 2B).
3. Deliver the impact.
   1. Identify the impact site at a point 2 mm left of the sagittal suture and 2 mm posterior to the coronal suture on the skull surface (Figure 2C).
   2. Align the impactor tip vertically over the target site, lower it gently until it contacts the skull surface, and ensure that the tip remains centered and perpendicular to the skull before releasing the weight.
   3. Release the 20 g weight from a height of 25 cm to deliver a single impact.
   4. For the Naive group, do not perform anesthesia or surgery. For the Sham group, administer anesthesia and make a scalp incision, but do not deliver the impact.
4. Suture the incision and provide postoperative care.
   1. Remove the impactor tip immediately after impact and inspect the surgical site.
   2. Close the scalp incision with interrupted 5-0 nonabsorbable polypropylene monofilament suture and disinfect the incision site with 10% povidone-iodine solution.
   3. Place the mouse on a thermostatic heating pad maintained at 37 °C ± 0.5 °C, monitor respiration until full recovery of consciousness and free ambulation, and then return the mouse to its home cage.
      NOTE: Recovery is defined by the return of regular spontaneous respiration, recovery of the righting reflex, and spontaneous ambulation. Acute apnea and visible hemorrhage should be monitored immediately after each impact and during early recovery. Animals that show prolonged apnea, visible hemorrhage, fail to regain the righting reflex within 10 min after discontinuation of isoflurane, or fail to ambulate spontaneously within 15 min should be excluded from further procedures and managed according to the approved animal protocol.
   4. Dispose of used blades and needles in an approved sharps container. Dispose of animal waste and contaminated materials in accordance with institutional biosafety regulations.
5. Repeat the injury.
   1. Repeat steps 3.1–3.4 every 24 h for 5 consecutive days to deliver a total of 5 impacts.
   2. Monitor body weight and activity continuously throughout the postoperative recovery period. Exclude animals from subsequent analyses if they develop a skull fracture, visible hemorrhage, seizure, persistent circling, or inability to feed or drink independently.

4. Acquiring magnetic resonance images

NOTE: In this study, MRI was performed on day 7 after the final injury. Mice were scanned under isoflurane anesthesia and were not sacrificed before imaging. Tissue collection was performed after imaging according to the experimental schedule.

1. Prepare the animal for MRI.
   1. Anesthetize the mouse with isoflurane and place it on an MRI-compatible animal bed equipped with a nose cone.
   2. Monitor respiration throughout the scan and maintain body temperature at 37 °C ± 0.5 °C.
2. Acquire T2-weighted images
   1. Acquire images on a 9.4 T superconducting magnet with a 30 cm bore and a gradient system with a maximum gradient strength of 1000 mT/m and a maximum slew rate of 10000 T/m/s.
   2. Use an 86 mm volume coil for radiofrequency transmission and a 15 mm planar surface coil for signal reception.
   3. Acquire T2-weighted high-resolution multi-slice fast spin-echo images using the following parameters: repetition time = 3000 ms, echo time = 46.34 ms, echo train length = 13, receiver bandwidth = 220 Hz/pixel, field of view = 20 × 20 mm², acquisition matrix = 208 × 208, slice thickness = 0.5 mm, slice gap = 0 mm, and number of excitations = 2.
   4. Acquire 30 slices to cover the entire brain. Use a total acquisition time of 1 min 33 s. In the present study, T2-weighted MRI was used for qualitative assessment only. No quantitative MRI analysis was performed.
      NOTE: MRI images were reviewed qualitatively by an investigator blinded to group assignment.

5. Assessing the gross neuropathology

1. Perform tissue collection.
   1. Deeply anesthetize the mouse according to the approved animal protocol.
   2. Open the thoracic cavity, expose the heart, insert a perfusion needle into the left ventricle, and incise the right atrium.
   3. Perfuse with normal saline, then perfuse with 4% paraformaldehyde (PFA). Perform the entire perfusion procedure in a certified chemical fume hood and avoid direct skin or eye contact with PFA. Dispose of tissues and liquid waste in compliance with hazardous waste and biosafety regulations.
   4. Collect the skull and intact brain and capture gross images using a high-resolution camera.
      NOTE: Gross pathological evaluation was performed by an investigator blinded to group assignment.

6. Monitor body weight

1. Record the body weight.
   1. Weigh each mouse before the first injury session and on days 1, 3, 7, 14, 21, and 30 after injury, and record the values to assess general health status over time.

7. Perform the beam-walk test

1. Prepare the apparatus.
   1. Use a horizontal wooden beam that is 1 m long and 1 cm wide and suspend it 30 cm above the ground.
2. Test motor coordination.
   1. Place the mouse at the starting end of the beam, allow it to traverse to the endpoint, and record the time required to cross the beam.
   2. Perform the test before injury and on days 1, 3, 7, 14, 21, and 30 after injury.
      NOTE: Behavioral assays were scheduled to minimize carryover effects between tests. Because the tail suspension test (TST) and forced swimming test (FST) are more stress-inducing than the open field test (OFT), they were performed first, followed by the OFT and Barnes maze at later time points. This sequence was selected to reduce potential confounding from prior habituation or repeated exploratory exposure during subsequent behavioral assessments.

8. Performing the tail suspension test

1. Prepare the apparatus.
   1. Perform the test on day 7 after injury and place a Plexiglas cylinder over the tail to prevent tail climbing. Use a cylinder that is 4 cm long with an outer diameter of 1.6 cm, an inner diameter of 1.2 cm, and a weight of 1.6 g.
2. Record the behavior.
   1. Fix the tail to the suspension box with adhesive tape, ensure that the mouse cannot reach the bottom of the box, and suspend the mouse for 3 min.
   2. Record the entire session with a high-resolution camera and analyze mobility time and immobility time manually from the recorded video by a blinded observer.

9. Performing the forced swimming test

1. Prepare the apparatus.
   1. Perform the test on day 10 after injury and fill a transparent cylinder 25 cm high and 15 cm in diameter with water maintained at 23–25 °C. Use a water depth that prevents the tail and hind limbs from touching the bottom.
2. Record the behavior.
   1. Place the mouse in the water for 6 min and record the entire session with a high-resolution camera.
   2. Analyze the final 4 min of the recording manually by a blinded observer and quantify mobility time and immobility time.
   3. Define immobility as only the minimal movements necessary to keep the head above water.

10. Performing the open field test

1. Prepare the apparatus.
   1. Perform the test on day 14 after injury in a quiet room under consistent illumination. This later time point was selected to provide separation from the preceding TST and FST sessions and thereby reduce acute stress-related carryover into the OFT.
   2. Use an open field box measuring 40 cm × 40 cm × 40 cm.
2. Record and analyze exploratory behavior.
   1. Place the mouse in the center of the arena and allow it to explore freely for 6 min.
   2. Track the behavior for 6 min using a behavioral tracking and analysis software. Calibrate the arena dimensions in the software to match the 40 cm × 40 cm apparatus and divide the arena virtually into a 5 × 5 grid.
   3. Define the 16 outer squares as the peripheral zone and the 9 inner squares as the center zone. Set the animal color darker than the background color. Track the animal’s body only; do not track the head or tail. Manually capture the background.
   4. Use the software to measure mean speed, time spent in the center zone, time spent in the peripheral zone, and total distance traveled over the full test period.
   5. Clean and deodorize the apparatus before and after each trial to remove olfactory cues.

11. Performing the Barnes maze

1. Prepare the apparatus.
   1. Begin the Barnes maze assessment on day 30 after injury.
   2. Use a circular stainless-steel platform 1 m in diameter with 20 evenly spaced holes.
   3. Place the escape box under one designated target hole, position visual cues around the maze, and keep the target hole location constant for all trials.
2. Conducting the training phase
   1. Turn on 90 dB white noise as an aversive stimulus during training.
   2. Place the mouse under a transparent cylinder in the center of the platform for 120 s.
   3. Remove the cylinder and allow the mouse to explore for up to 180 s.
   4. Record a successful trial when all four paws enter the escape box.
   5. Guide the mouse into the escape box if it fails to locate the target within 180 s and record the latency as 180 s.
   6. Allow the mouse to remain in the escape box for 60 s.
   7. Repeat the training trial once after a 4–5 h interval on each training day and continue training for 4 consecutive days.
3. Conducting the rest day and test phase
   1. Suspend training for 1 day after the training phase.
   2. Place the mouse in the center of the platform on the test day and allow exploration for up to 180 s.
   3. Set the maze diameter to 100 cm for scale calibration, and ensure that the tracking arena is centered in the video frame. Set the animal color darker than the background. Track the animal’s body only; do not track the head or tail. Manually capture the background.
   4. Record the latency to enter the escape box, mean speed, and total distance traveled during the 180 s test session using a behavioral tracking and analysis software. Define the escape zone in the software according to the location of the designated target hole before analysis.

12. Data analysis

1. Prepare the dataset.
   1. Compile the behavioral, imaging, and body weight data in a statistical and graphing software.
   2. Express the data as the mean ± standard error of the mean (SEM).
2. Perform statistical analysis.
   1. Analyze longitudinal body weight and beam-walk data in this study using a linear mixed-effects model with group, time, and their interaction as fixed effects and animal as a random effect, followed by post hoc multiple-comparison tests when applicable.
   2. Analyze TST, FST, OFT, and Barnes maze outcomes using ordinary one-way ANOVA for normally distributed data with homogeneous variances, followed by Tukey’s post hoc test when the overall ANOVA was significant. For non-parametric data, use the Kruskal-Wallis test followed by Dunn’s post hoc test.
   3. Report exact p.-values whenever available, together with the test statistic, degrees of freedom, and an effect size measure (R² for one-way ANOVA). Derive all reported p-values from the statistical tests specified above and perform all analyses in the statistical and graphing software.
   4. Set statistical significance at p. < 0.05.

Results

Imaging and neuropathological evaluation of c-TBI mice

To assess the overall health status and basal motor function of the animals, body weights and beam-walk performances were monitored according to the timeline shown in Figure 1. Longitudinal group differences were analyzed using a linear mixed-effects model with animal as a random effect. Prior to modeling, there were no statistical differences in body weight among the groups (

Discussion

This study details a standardized experimental protocol for establishing a repetitive mild traumatic brain injury (rmTBI) mouse model using a repeated free-falling weight-drop method. By securing the skull-exposed mouse in a stereotaxic apparatus and applying repeated impacts to defined coordinates using a 20 g weight dropped from a height of 25 cm for five consecutive days, a behavioral phenotype characterized by chronic depressive-like behavior was induced without skull fracture or gross focal brain contusion. The prin...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
5-0 nonabsorbable polypropylene monofilament suture	Jinhuan Medical	I303	wound closure
Anesthesia machine	RWD Life Science	R550	construction of animal models
ANY-maze behavioral tracking software	Stoelting	Version 7	behavioral software
Barnes maze	Shanghai Xinruan Information Technology	XR-XB108	behavioral software
Biorender	BioRender	https://biorender.com	scientific research drawing software
Buprenorphine	Sigma-Aldrich	Y0001108	postoperative analgesia
Electric shaver	AUX	AUX-C5-1	preoperative hair removal
Forced swimming test	Shanghai Xinruan Information Technology	XR-XQX201	behavioral software
GraphPad Prism 9.5	La Jolla	9.5	statistical software
Isoflurane	RWD Life Science	R510-22-10	construction of animal models
Mice	Charles river	C57/6N	experimental animal
Open field test	Shanghai Xinruan Information Technology	XR-XZ301	behavioral software
Ophthalmic ointment	BEIJING TWINLUCK PHARMACEUTICAL	H11021342	prevent corneal drying during anesthesia
Paraformaldehyde (PFA), 4%	Biosharp	BL539A	perfusion and tissue fixation
Phosphate-buffered saline (PBS)	Biosharp	BL1425A	perfusion and tissue washing
Povidone-iodine solution, 10%	Beyotime	27883-25g	skin disinfection before surgery
Preclinical Magnetic Resonance Imaging System	United Imaging Life Science Instrument	uMR9.4T	imaging software
Scalpel blade No. 11	Jinhuan Medical	K3-11	skin incision during surgery
Scalpel handle	Jinhuan Medical	K6-10	used with No. 11 scalpel blade
Stereotaxis instrument	RWD Life Science	68025	construction of animal models
Sterile gauze	HYNAUT	BK05	bleeding control and surgical cleaning
Tail suspension test	Shanghai Xinruan Information Technology	XR-XQX201	behavioral software
Weight-drop impactor	RWD Life Science	68093	construction of animal models