JSM received funding from the United States Department of Veterans Affairs (VA) Office of Biomedical Laboratory Research &#38; Development (JSM, I01BX004896) and the VA Northwest Mental Illness Research Education and Clinical Center, a Congressionally-mandated VA entity investigating blast-induced brain injury and comorbid post-traumatic stress. JSM reports unrelated funding from the FY22 Traumatic Brain Injury and Psychological Health Research Program Translational Research Award (W81XWH-22-TBIPHRPTRA, Award Number HT94252310755). The authors thank Andrew Shutes-David for his editorial assistance.

All procedures were performed under protocol #1588223, approved by the Veterans Affairs Puget Sound Health Care System Institutional Animal Care and Use Committee and in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals.
1. Animal care
NOTE: Animal models of LLB are limited solely by their availability and the capacity of the shocktube to accommodate their size. The described shocktube herein was designed specifically for use with mice.
<ol>
	<li>Use 3-4-month-old male or female C57BL/6J mice or other approved mouse strains/lines in accordance with experimental needs. Maintain the mice on a 12 h dark-light cycle in specific pathogen-free facilities with ad libitum access to food and water. Mice are typically socially housed with 4 or 5 in a cage. Maintain facility temperatures at 20-22 &#176;C.</li>
	<li>Bring cages containing blast and sham mice to a nearby holding area. Bring separate empty cages for transferring individual mice to and from the blast room.</li>
</ol>
2. Shocktube preparation
<ol>
	<li>(Safety check) Confirm that the necessary safety checks have been completed for the specific system. Ensure that the gas supply (helium) and master power are off/disconnected.</li>
	<li>Prepare membranes as needed for the specific number of low-intensity blasts to be conducted (Figure 1.1). Cut membrane dimensions as required for the specific shocktube used in this protocol:
	<ol>
		<li>Cut one sheet of plastic cling wrap into a 5.5&#34; x 5.5&#34; square to seal the spool, allowing it to pressurize.</li>
		<li>Cut one sheet of standard 8.5&#34; x 11&#34; copier paper (75 g/m2 weight) to 5.5&#34; x 11&#34;; fold the resulting sheet of paper in half to form a 5.5&#34; x 5.5&#34; square.</li>
		<li>Obtain one sheet of 500 G mylar membrane (125 &#181;m thickness). 
		NOTE: These sheets are not ruptured or significantly deformed by standard low-intensity blasting and can be reused for the duration of a day&#39;s procedure.</li>
	</ol>
	</li>
	<li>Take a square of cling wrap and a square of folded paper and set them out on a flat surface (Figure 1.2). Place the folded paper on top of the cling wrap and align the two with each other as best as possible (Figure 1.3). To expedite repetitive blasts, arrange all the membrane stacks now.</li>
	<li>Insert the mylar membrane between the driver and the spool by rolling it up into a small tube (about as big around as one&#39;s index finger; Figure 1.4,1.5). Insert it completely into the mechanism, and let go to allow it to unroll against the rubber seal that separates the driver section from the spool. Push the spool toward the driver to secure the mylar sheet in place; this will unseal the spool from the driven section of the shocktube.</li>
	<li>Place the fingers under the top half of the cling wrap and carefully roll both the cling wrap and paper toward you, ensuring they roll up together without becoming misaligned (Figure 1.6).</li>
	<li>Insert the membrane stack between the spool and the driven sections of the shocktube (Figure 1.7).</li>
	<li>Allow the membrane stack to unroll so that the plastic seal is facing toward the spool and the paper is facing toward the driven section of the tube (Figure 1.8). 
	NOTE: This orientation will create an airtight seal so that the system can be pressurized.</li>
	<li>Close the spool assembly (Figure 1.9,1.10). As appropriate, tighten the bolts by hand or hydraulically, securing the driver-spool-shocktube assembly so that the system can be pressurized. (Safety check;&#160;Figure 1.10) 
	NOTE: For hydraulic systems, ensure that the closure assembly&#39;s target pressure is reached to prevent misfires, which can require membrane replacement and slow the LLB exposure process. We use hydraulics to close our assembly at 500 psi.</li>
</ol>
3. Animal preparation
<ol>
	<li>Turn on the circulating water heating pad below the anesthesia chamber, with the temperature set to 37 &#176;C (Figure 1.11). Place an absorbent medical pad on top of the heat pad.</li>
	<li>In the holding room, remove one mouse from its home cage and place it into an empty transfer cage. Bring the caged mouse into the blast room.</li>
	<li>Turn the oxygen flow rate to 1.0 L/min (lpm) and the vacuum scavenge system on (Figure 1.12).</li>
	<li>Turn the isoflurane on to 5% (to induce rapid unconsciousness) and route the flow to the rodent anesthesia chamber (Figure 1.13).</li>
	<li>Place the mouse in the chamber to induce anesthesia (Figure 1.14).</li>
	<li>Once the mouse is fully anesthetized and displays stable breathing for an additional 30 s, reach into the chamber and ear punch the mouse for unambiguous long-term identification of the mouse throughout the remainder of the study. Doing this step now is necessary to avoid interfering with recovery times after the blast. Then, apply sterile ophthalmic lubricant to both eyes to prevent corneal drying.</li>
	<li>Remove the mouse from the chamber and place its nose into the nosecone (Figure 1.15). Switch the flow of anesthesia (e.g., isoflurane) from the induction chamber to the nosecone.</li>
	<li>Use small pieces of laboratory tape to lightly restrain the mouse&#39;s limbs against the gurney (Figure 1.16).</li>
	<li>After restraining the mouse, place a wire twist tie around each limb and twist tightly, securing the mouse to the gurney at the wrists and ankles (Figure 1.17). Place a larger twist tie around the chest, tying it very loosely such that the mouse&#39;s breathing is not restricted. This will serve as a secondary restraint mechanism in case any of the limb restraints come loose.</li>
	<li>Lift the mouse&#39;s tail and place it under the left foot to ensure it does not get pinched when the gurney is inserted into the shocktube (Figure 1.18).</li>
</ol>
4. LLB procedure
<ol>
	<li>Open the animal exposure section of the shocktube and orient the mouse so that it faces the oncoming blast wave (Figure 1.19).</li>
	<li>Secure/suspend the gurney in the animal exposure section (Figure 1.20).</li>
	<li>Close the door tightly for the animal exposure section, ensuring that the anesthetic flow tube is not pinched by the door (Figure 1.21).</li>
	<li>Reduce the anesthesia to 2.5-3% isoflurane, 1 lpm for the remainder of the session.</li>
	<li>Power the system as appropriate (Figure 1.22).</li>
	<li>Locate and connect the supply line for the compressed helium gas (Figure 1.23,1.24).</li>
	<li>Leave the blast room to access the blast tube control console in an adjoining room, and ensure no personnel or animals are left in the blast room. 
	NOTE: Hearing protection may be required by the institution or by operational conditions. Such conditions may include shocktube arrangements where the control console is located in the same open space as the shocktube.</li>
	<li>From the console, turn on the acquisition software to record the blast event (see the green box in Figure 1.25). 
	NOTE: For these procedures, we collect sensor data at a 20 kilo hertz (kHz) sampling rate, which is then processed using LabView software. We recommend acquiring sensor sampling at &#8805;10 kHz to achieve high-quality time versus pressure curves.</li>
	<li>Disengage any safety lock (e.g., power control keys, which are depicted by a green arrow in Figure 1.26).</li>
	<li>Close both gas vents and passively pressurize the spool (Figure 1.27). Do not use the driver side. Continue to fill until the membrane ruptures on its own at the target peak psi as determined by the number of membrane sheets used.</li>
	<li>Record the peak pressure, positive phase duration, and impulse at the animal location. (Figure 1.28). Turn off the fill mechanism.</li>
	<li>Return to the shocktube, disconnect the helium feed line, and turn off the power supply to the blast control circuit (Figure 1.29).</li>
	<li>To conduct repeated LLB exposures on the same animal, open the spool, remove the spool membrane stack, and then roll and insert another spool membrane stack (Figure 1.30, 1.31, 1.32). Flatten the membrane stack and reclose the assembly. 
	NOTE: To model the clinical experience of low-level blast exposures during empirically defined SOF training, we expose mice to 5-6 LLBs per day, capping daily exposures to a conservative ~20 cumulative total psi45. Studies emphasizing mechanistic and dose-response relationships may alternatively choose to use a consistent number of LLB exposures with defined overpressure parameters per session.</li>
	<li>After the final LLB for the current animal, remove it from the shocktube, leaving the anesthesia on (Figure 1.33).</li>
	<li>Untie the animal while it is under anesthesia. Remove it from the anesthesia nosecone, placing it on its back onto the heated water pad (Figure 1.34).</li>
	<li>Once the animal has been placed on the water pad, start a timer and record the amount of time until the mouse flips over onto its ventral side (i.e., its stomach) on its own (Figure 1.35). Record this time as the righting time. Once the mouse recovers, return it to the home cage and continue monitoring as needed.</li>
</ol>
5. Multiday procedures
<ol>
	<li>To model routine LLB exposures from breaching charges used during SOF Close Quarter Battle training, perform repeated daily exposures on the mice 5 days a week (Monday through Friday) for a total of 15 days across 3 standard work weeks.</li>
</ol>
6. Altering peak LLB pressures
<ol>
	<li>Increase peak pressure through the use of stronger membrane materials or by simply stacking additional membranes. For example, use Mylar Roll Clear 0.005 (500 G) membrane to produce ~20 psi peak pressure (when used as both driver and spool membranes) or Mylar Roll Clear 0.002 (200 G) membrane to produce ~10 psi peak pressure.</li>
	<li>Adjust the parameters for the positive phase duration and impulse of the blast to meet experimental needs. To adjust positive phase durations and impulses, empirically determine target conditions by substituting compressed gas sources47,49 or changing the driver length whenever possible. The above protocol uses helium to create a sharp peak pressure and waveform similar to an idealized Friedlander curve.</li>
</ol>
7. Tissue collection
NOTE: Tissue collection practices can be adjusted according to experimental needs.
<ol>
	<li>Anesthetize the mouse via intraperitoneal injection with 210 mg/kg of pentobarbital.</li>
	<li>Place the mouse into a mouse or rat cage with bars or a premade mesh; place the caged mouse into a fume hood.</li>
	<li>Once the mouse is unresponsive, place it on its back on the bars on top of the cage and close its mouth around one of the bars to help it stay in place during perfusion.</li>
	<li>Grab the skin of the stomach, pull it upwards, and use a pair of large scissors to cut a hole in the abdominal cavity, being careful not to cut any of the organs. Continue to cut farther down along the base of the ribs to allow for freer articulation of the ribcage.</li>
	<li>Using a hemostat, approach the mouse from the side and grab the tissue directly on top of the ribcage, rolling the hemostat back to keep the base of the ribcage angled in an easily accessible position. Use a pair of forceps or a similar tool to hold the hemostat in place.</li>
	<li>Using a small pair of surgical scissors, carefully cut the diaphragm to allow access to the heart. Use a pair of forceps to gently angle the heart such that the bottom is facing directly out of the open base of the ribcage. Work quickly so the heart will still be beating during perfusion.</li>
	<li>If collecting blood, hold the heart with a pair of forceps and carefully pierce the right ventricle using a 3 mL syringe tipped with an 0.5&#34; 25 G needle. Insert from the bottom of the ventricle and go in lengthwise, being careful not to pierce the opposite side of the ventricle. Gently pull on the syringe until 0.5-1.0 mL of blood has been collected or the flow stops, and then remove the syringe.</li>
	<li>Use a pair of surgical scissors to cut a small incision in the right atrium to allow blood and perfusate to drain. Hold the heart with a pair of forceps, and carefully insert a 25 G butterfly needle into the left ventricle, inserting from the bottom. Hold the butterfly needle in place with a holding clamp or by hand.</li>
	<li>Perfuse the animal.
	<ol>
		<li>Connect a syringe containing 50 mL of phosphate-buffered saline (PBS) to a butterfly needle and perfuse at a rate of approximately 10 mL/min. Look for blanching of the liver as a sign of proper perfusion. After the syringe is emptied, disconnect it from the butterfly needle.</li>
		<li>For preparation of tissues for microscopy, replace the empty PBS syringe with a syringe containing 50 mL of 10% neutral buffered formalin (NBF) or 4% formaldehyde solution. Repeat the above steps to perfuse with formalin. 
		NOTE: The perfused mouse should be observed to twitch during perfusion; this should result in full body rigor or rigidity after the procedure is completed.</li>
	</ol>
	</li>
	<li>Remove the butterfly needle from the heart and remove the mouse from the cage bars for tissue collection.</li>
	<li>Remove and subdissect the target organs according to need; be careful to perform procedures on ice when fresh, unfixed materials are collected.</li>
	<li>Flash freeze any unfixed tissues that were collected in liquid nitrogen and store them at -80 &#176;C until used in protocols assaying protein or RNA targets.</li>
	<li>For fixed tissues, remove to a labeled 50 mL conical tube filled with formalin (one tube per organ).</li>
</ol>

Shocktube Method to Model Highly Repetitive Low-Level Blast in Mice

The authors have no financial interests to disclose.

We cannot adequately treat what we inadequately understand, and we do not yet understand the injury mechanisms related to highly repetitive LLB exposure. Many SOF personnel report the development of health-related impairments thought to be related to highly repetitive LLB exposure within five to ten years of operational service50,51. Some SOF personnel develop acute traumatic brain injury (TBI)-like neurocognitive effects immediately after LLB exposure39. Furthermore, clinicians report that symptoms resulting from blast exposure are frequently refractory to traditional treatments, which may drive SOF and clinicians to look to alternate treatments52,53. Despite the frequent exposure of SOF to LLB and overpressure mechanisms45, the severity and treatment resistance of the resulting symptoms, and the documented pattern of blast-related astroglial scarring51, the long-term health outcomes remain relatively unknown. Clinicians and military leadership rely on modeling research to uncover injury mechanisms and pathophysiology. These models are critical to developing policies and strategies to identify, interrupt, prevent, and treat the pathology process early.
Crucially, mouse modeling of common military LLB exposures is expected to inform health-prediction models. Clinical practice would benefit from LLB predictive models that identify who may be at greatest risk for blast-related pathology, which blast properties provoke the most serious outcomes, and how the disease process may evolve based on the chronicity, dosing, or specificity of the blast exposure. Thus, modeling repetitive LLB exposure is essential in developing hypotheses and predictions for how exposures will impact the health outcomes of SOF and other Service members. Prediction and injury mechanism models would inform diagnostics and treatment, as well as return-to-duty decisions based on symptoms and exposure.
The study of blast-induced TBI (bTBI) in mice has seen significant advancements in recent years, particularly with the development of models that predict outcomes following chronic repetitive mild bTBI in humans54,55. Whereas the study of mid-to-high level blast exposure using shocktubes is well developed with hundreds of PubMed indexed articles46,56,57,58, shocktube use in studies of blasts near routine military training overpressures (&#60;6 psi peak pressure40) is less developed, with fewer than ten articles identified in a recent PubMed search19,20,22,23,26,27,28. To facilitate the development of this understudied field, the presented model focuses on key considerations for consistent LLB overpressures in mice, post blast recovery, and monitoring while noting several distinct advantages of this model over the use of open-field explosives. Indeed, we argue that the described laboratory LLB model may enable the development of predictive models of clinical outcomes following chronic repetitive LLB.
The LLB model offers critical advantages over open-field explosive blast models, particularly in terms of animal welfare. Open-field models may result in 3-8% mortality rates25,36, whereas this lab-based LLB model shows no loss. This distinction is crucial, especially when simulating the high cumulative exposures typical of military training, where virtually no trainees experience fatal outcomes from LLB exposure. The apparent absence of apnea or other causes of death, such as lethal pulmonary trauma, ensures the model&#39;s reliability and consistency, positioning it as a preferred choice for studies on the clinically relevant effects of repetitive LLB.
This protocol is specific for an &#34;open-ended&#34; shocktube with a three-part design, consisting of driver, spool, and driven sections. Highly repetitive LLBs may be achievable with other shocktube designs with appropriate modifications to the protocol. Open-ended shocktube designs are frequently used for the study of blast-induced neurotrauma46,47,48. The open-ended shock tube, featuring an open exit end, allows the generated shock wave to propagate freely down the length of the tube where it encounters its target (e.g., the animal subject) before exiting the opposite end of the tube. This design facilitates the reproduction and study of relatively pure primary blast overpressures approximating the properties of blast explosions as they would occur in the open field48. As a result, the fidelity of the empirically measured blast overpressure wave is compared with an idealized Friedlander wave; this allows the evaluation of the tube performance to produce a specific overpressure event. To model LLB exposure, we use a previously described48, custom-built, open-ended blast tube originally designed to reproduce the effects of HLB detonations of over 200+ lbs of trinitrotoluene (TNT) at a standoff distance of ~25 feet. To enable high peak overpressures, a gas is pressurized into the driver, which is separated from the spool by a membrane, sealing the gas in the driver. The spool, in turn, is also separated from the open-ended section by another membrane. This second membrane allows the spool to be separately pressurized. The dual chamber system allows the gases in the driver to be pressurized past the membrane&#39;s normal point of rupture. This occurs because the pressurized spool acts as a buffer, supporting the membrane at the interface of the driver and the spool, thus preventing its rupture. When the shock tube operator desires to generate a shockwave at the target pressure, an electronic valve vents gas from the spool, rapidly dropping pressure in the spool and allowing the overpressurized gas in the driver section to rupture both the driver and spool membranes and rapidly expand down the length of the tube where it encounters the animal in the target zone. The key modification enabling the study of LLB in high-performance tubes of this design is that we block off the driver and only use the spool in combination with low-threshold membranes.
To ensure the reliability and reproducibility of LLB experiments, certain actions must be taken during the setup. Tightly securing the arms and legs at the wrists and ankles is crucial. This minimizes variability in bodily movement and blast exposure and prevents unintended injuries that could confound results. Additionally, rotating wrists and ankles inward helps direct the movement of the appendages toward the animal midline, reducing the risk of distal injuries that might affect subsequent motor performance assessments. The straightening of the head and spinal curvature is another essential factor in ensuring uniform blast exposure across subjects, as it helps to reduce potential differences in range of motion. Increasing the percentage of isoflurane that is used for anesthesia is recommended for protocols spanning multiple days or weeks. This adjustment helps maintain consistent anesthesia depth throughout the extended experimental duration. In our experience, an increase in 0.5% isoflurane is sufficient to maintain adequate anesthesia.
However, anesthesia delivery via nosecone may not be possible for all blast tube designs, especially for those with full enclosures that do not allow the insertion of the tubing into the driven section. In such events, injectable anesthetics may be preferable. We recommend determining how much time is required for the delivery of the repeated sequential blasts and then administering sufficient anesthetic to maintain unconsciousness throughout the procedure. Additional animal welfare checks may be necessary during the development of this modified method to ensure proper anesthesia maintenance. Furthermore, the use of injectables may render postacute response monitoring, such as the collection of righting time measures, impossible.
Ethical considerations are paramount in animal research, and this lab-based LLB model incorporates comprehensive post-blast recovery and monitoring protocols. Humane endpoints after blast exposure, including difficulty breathing, inability to right themselves, non-ambulatory status after a 2 h observation period, seizure-like movements, awkward movements, vision impairment, and evidence of internal bleeding or fractured limbs, are closely observed. Notably, LLB blast mice have not exhibited any of these conditions in our experiments. However, limb fractures can occur during HLBs, often due to operator error. Mitigating this risk involves rotating the hands and feet toward the animal midline during gurney securing. This technique prevents the blast wind from sweeping the appendages backward and breaking the associated bones.
This repetitive LLB model&#39;s advantages extend beyond ethical considerations to practical and methodological aspects. Its lab-based design eliminates the need for handling explosives, thereby enhancing safety and accessibility. The model is highly reproducible and customizable, allowing researchers to influence exposure parameters through the use of different gas types, device settings, and membrane strengths. Helium, which is chosen here for its ability to reproduce open-field explosion kinetics49, may provide a reliable baseline47,59,60. Adjusting peak pressure is achieved empirically by modifying the retention membrane thickness or strength, enabling fine-tuning for specific experimental requirements. Lastly, the LLB model eliminates the impact of seasonal or weather variations on data, animal exposure, and other experimental factors. This consistency ensures robust and reliable results, making this repetitive LLB model an invaluable tool for longitudinal and highly repetitive blast research.
Understanding blast-related neurotrauma requires elucidating injury mechanisms, blast intensity metrics, and threshold values. However, uncertainties surround human brain injury mechanisms in blast scenarios. Previously proposed criteria for human injury following blast exposure have relied on animal studies, yet it is challenging to directly apply these studies to humans due to incomplete scaling criteria across species61. Lung injury scaling based on animal body mass is an exception, given the presence of accepted criteria62,63. Proposed scaling laws for brain effects, based on body64,65 or brain mass66, however, overlook known and unknown anatomical differences, especially concerning the protective structures inside and around the brain. Mass scaling predicts higher injury risks in smaller-bodied species, which are contradicted by studies both in birds67,68,69 and humans70. Developing accurate scaling laws therefore demands an empirical understanding of the relationship between external blast event intensity and internal brain effects across species. In the case of LLBs, very little is known about either single or chronic exposure in either animal models or people. As a result, the empirical studies needed to inform the development of future scaling laws at the LLB intensity range may be catalyzed by our method.
In summary, this laboratory-based shocktube model represents a significant advancement in the study of the chronic effects of LLB exposure in mice. By incorporating procedures for modeling consistent overpressures, prioritizing post-blast recovery and monitoring, and highlighting distinct advantages over alternative models, this lab-based LLB model may provide a reliable and ethical choice for advancing our understanding of injuries related to chronic LLB exposure.

Low-level blast (LLB) exposure occurs when individuals or structures experience relatively low magnitudes of explosive force, typically arising from small industrial accidents, controlled demolitions, or certain military training activities. In contrast, high-level blast (HLB) exposure entails exposure to intense and potentially destructive magnitudes of explosive force, commonly encountered in military combat, terrorist attacks, or large-scale accidental explosions. The primary distinction between LLB and HLB therefore lies in the intensity of the explosive events and, by extension, the ability of exposed persons to tolerate repeated exposures before experiencing physical or functional injury. In this regard, the effects of HLB exposure tend to be more obvious than the effects of LLB exposure. Because of this, persons with significant LLB exposure may be at increased risk for slowly developing injuries or deficits that go undetected until their cumulative effects become discernable.
Ongoing research aims to enhance our understanding of how the properties of blast exposure, such as intensity or repetition, may cause injury so that we can better guide prevention and medical management. In military medicine, understanding the clinical implications of blast exposure is of paramount importance, and as a result, animal models capable of informing those outcomes are needed. Although animal models have helped elucidate the effects of HLB, the effects of LLB exposures remain largely understudied. Numerous modeling studies examine the effects of blast overpressures near or above 10 pounds per square inch (psi) peak pressure1,2,3,4,5,6,7,8,9,10,11,12,13,14,15,16,17,18, but few reports focus on pressure levels ranging from 1 to 7 psi19,20,21,22,23,24,25,26,27,28,29,30,31,32,33,34,35,36, which are more common in military training environments37,38,39,40 and fall near the historical threshold of 4 psi for safe environmental exposure. Thus, broader dissemination of methods for the study of frequently used peak pressures of LLB may help catalyze rapid clinical insights for application to military medicine and force optimization.
A significant association between the occupational risk of LLB and diverse clinical diagnoses is emerging from epidemiological investigations of military LLB41,42,43,44. These studies support a poorly defined dose-dependent relationship, with repetitive LLB exposures demonstrating heightened risks41. This suggests that increasing cumulative blast exposure plays a crucial role in shaping clinical outcomes in military settings.
Previous animal modeling studies of LLB under 10 psi have primarily used explosives or shocktube systems to investigate the effects of exposure. Although these models typically examine the effects of one to three exposures, they have nonetheless contributed to a growing understanding of the mechanistic19,20,30,31, neuropathological29,31,33, and behavioral consequences19,20,23,25,32,34, associated with low-intensity blast exposures that are typical of the military training environment.
Studies examining single LLBs generated by open-field explosives have reported evidence of subtle brain pathologies and behavioral changes frequently associated with posttraumatic stress. Woods and colleagues24 were unable to detect microscopic brain injury at 2.5-5.5 psi, but they did detect quantitative changes in brain tissue glycosphingolipids by mass spectrometry. Using the same peak pressures and experimental design, Rubovitch and colleagues25 observed behavioral changes following blasts that occurred with a similar lack of brain pathology when measured by light microscopy. However, in subsequent pathological investigation, unambiguous ultrastructural damage to brain myelin, mitochondria, neurons, and neurovasculature was identified by electron microscopy29,30,31,32,33 in 6.7 psi LLB-exposed mice. Interestingly, several LLB studies using open-field explosives with pressures of ~10 psi and less report approximately 3-8% mortality after a single exposure25,36.
Similar results have been previously noted by several studies using laboratory shocktubes. In studies examining single LLBs produced by shocktubes, evidence has been found of neural cytoskeletal injury and changes in neuronal firing patterns developed after exposure to a single 1.7 psi blast22. At 4 psi, corpus callosum dysfunction was reported to accompany neurobehavioral deficits in LLB-exposed rats23. Compared to the blast duration measured in air, Chavko and colleagues27 found that the positive phase duration of the blast overpressure was significantly longer in the brains of rats exposed to 5.8 psi. Biosignatures of similar injury responses may be supported by a study in mice following 7.5 psi exposure in which Ahmed and colleagues35 report detectable changes in serum levels of specific inflammatory, metabolic, vascular, and neural injury proteins up to a month after exposure. Interestingly, this study also reported 4.5% mortality at 24 h following exposure.
In studies examining three shocktube LLBs over a single 20 min exposure session, LLBs between 1.4 and 8.7 psi caused psi-dependent increases in intracranial pressure (ICP) in rats, with observable ICP changes taking longer for lower psi20 and resulting in cognitive changes19,20. Using pigs, the same group determined that three 4 psi LLB exposures from a variety of military equipment were sufficient to cause histological neuropathology when the animals were placed in gunner positions simulating human use of the equipment21.
These studies collectively illustrate the diverse effects of LLB exposure that may occur under conditions of limited exposure and recovery periods. Repetitive LLB exposure appears to induce persistent cognitive and behavioral deficits, emphasizing the need for a nuanced understanding of the cumulative effects so we can better determine when those effects may become clinically significant; this is particularly relevant for military trainees who are exposed to high levels of repetitive LLB. To achieve this, new studies are required since the current literature does not adequately model the clinical experiences of routine military training exposures that exceed one to a few blasts over the course of a few days.
Special Operations Forces (SOF) may endure significant and highly repetitive LLB during routine exposures. A recent study estimates the representative exposure anonymized across all positions in an explosive entry breaching team to be as high as 184 cumulative peak psi over the course of one training week42. This is based, in part, on a conservative estimate of 6 breaching charges used per day, with an average of 4 psi peak pressure each, as measured by personnel-mounted blast gauges; it does not account for flashbangs and other devices45. A routine training cycle may last several weeks. To facilitate the study of clinical LLB experiences, such as those of training SOF members, we present a laboratory-shocktube model of highly repetitive LLB exposure. The method, based on established pneumatic shocktube systems46,47,48, allows for highly reproducible investigations of pressures of 2 psi and higher. The procedure is not dependent on external factors such as weather, results in no observed mortality, and is lab-based. As a result, the method enables sustained, daily repetitive LLB exposures in the same subjects for studies lasting weeks to months, facilitating the high-fidelity investigation of military training.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Adroit Thermal Recirculating Heat Pump (120 V)</td>
			<td>Parkland Scientific</td>
			<td>HTP-1500</td>
			<td></td>
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			<td>Copy paper, 75 g/m2 weight</td>
			<td>Staples</td>
			<td>897804</td>
			<td></td>
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			<td>Disposable Absorbant Blue Pads</td>
			<td>VWR</td>
			<td>82020-845</td>
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			<td>Forane Inhalant Solution</td>
			<td>MedLine</td>
			<td>10019-360-60</td>
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			<td>Helium</td>
			<td>Linde</td>
			<td>UN1046</td>
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			<td>Laboratory tape (1&#34;)</td>
			<td>VWR</td>
			<td>89098-076</td>
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			<td>LabView software</td>
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			<td>Medical oxygen</td>
			<td>Central Welding Supply</td>
			<td>UN1072</td>
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			<td>Mylar, 0.005 thickness</td>
			<td>Tapp Plastics</td>
			<td>22934</td>
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			<td>Plastic cling wrap</td>
			<td>Santa Cruz Biotechnology</td>
			<td>sc-3687</td>
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			<td>Plastic twist ties&#160;</td>
			<td>VWR</td>
			<td>11215-940</td>
			<td></td>
		</tr>
		<tr>
			<td>Pneumatic Shocktube (with driver and spool sections; target area sized for mice, 20 kHz sampling rate pressure sensors, control and acquisition software)</td>
			<td>BakerRisk, San Antonio, TX</td>
			<td>custom</td>
			<td></td>
		</tr>
		<tr>
			<td>Reusable Heavy Duty Heating Pad (12&#34; x 18&#34;)</td>
			<td>Parkland Scientific</td>
			<td>121218</td>
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modeling highly repetitive low-level blast exposure in mice

Exposure to explosive blasts is a significant risk factor for brain trauma among exposed persons. Although the effects of large blasts on the brain are well understood, the effects of smaller blasts such as those that occur during military training are less understood. This small, low-level blast exposure also varies highly according to military occupation and training tempo, with some units experiencing few exposures over the course of several years whereas others experience hundreds within a few weeks. Animal models are an important tool in identifying both the injury mechanisms and long-term clinical health risks following low-level blast exposure. Models capable of recapitulating this wide range of exposures are necessary to inform acute and chronic injury outcomes across these disparate risk profiles.
Although outcomes following a few low-level blast exposures are easily modeled for mechanistic study, chronic exposures that occur over a career may be better modeled by blast injury paradigms with repeated exposures that occur frequently over weeks and months. Shown here are methods for modeling highly repetitive low-level blast exposure in mice. The procedures are based on established and widely used pneumatic shocktube models of open-field blast exposure that can be scaled to adjust the overpressure parameters and the number or interval of the exposures. These methods can then be used to either enable mechanistic investigations or recapitulate the routine blast exposures of clinical groups under study.

While investigating experimental outcomes in mice following exposures to explosive blast forces, recording and characterizing the event through pressure versus time analysis is crucial for evaluating the success of the experiment. This method, which involves measuring the dynamic changes in pressure during the blast, helps investigators understand the effects of blasts on biological systems.
In successful experiments, pressure recordings exhibit a well-defined and controlled wave pattern. The pressure rise is sharp, reaching peak values within expected times (Figure 2). The subsequent pressure decline follows a predictable curve, exemplified by the Friedlander waveform, indicating efficient dissipation of energy. In terms of injury assessment, no overt signs of injury are present in LLB experiments, even when conducting highly repetitive LLB exposure with up to six blasts occurring within 15-20 min (Figure 3). However, an analysis of righting times following repetitive LLB exposure indicates that blast mice return to consciousness faster than sham mice (Figure 4). Thus, repetitive LLB results in reproducible changes in acute neurobehavioral arousal responses after exposure.
Suboptimal experiments may display irregular pressure profiles. Instances in which peak pressures are unexpectedly depressed may indicate a premature or slow release of gas, preventing the sharp release of gas expansion down the length of the driven shocktube section to encounter the animal in the target area. Premature loss of gas pressure is often the result of improperly sealed driver or spool sections. This can result from flaws in the membrane or inadequate tightening of the driver-spool-shocktube assembly. In such cases, biological samples may exhibit reduced signs of trauma.
Data interpretation involves linking pressure-time profiles with observed biological responses. Successful experiments demonstrate that the chosen blast parameters, such as peak pressure and duration, elicit the expected or established biological responses under investigation. Correlations between specific pressure features and biological outcomes aid in establishing causal relationships. Longitudinal studies are enabled by this protocol due to the lack of observed animal loss for study time points as long as 6 months after the final LLB (Figure 5).
The range of clinical outcomes following LLB exposure is subtle and poorly understood. Repetitive exposure to LLBs has historically been considered subinjurious for both people and mice. This is supported by a quick return to normal ambulation, behavior, and physical activity following exposures at 2-5 psi. However, the lack of overwhelming acute neurosensory symptoms or behavioral changes does not preclude the existence of negative insidious effects. Because LLB-related phenotypes are subtle at best, the full range of effects is an area of active investigation and may require considerable time or repetition to provoke clinically significant outcomes.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66592/66592fig01.jpg" /> 
Figure 1: Procedural steps for the shocktube model of repeated murine LLB. Following both the preparation of the shocktube (Steps 1-10) and the animal preparation stages (Steps 11-18), mice are exposed to one or more LLBs (Steps 19-32), before being removed from the tube (Step 33). Mice are then placed on their backside onto a warmed heating pad (Step 34). The amount of time it takes the animal to flip over onto their ventral side is recorded as the righting time (Step 35). Abbreviation: LLB = Low-level blast. <a href="https://www.jove.com/files/ftp_upload/66592/66592fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66592/66592fig02.jpg" /> 
Figure 2: Representative pressure-time curves for exposures near 4 psi. (A) Additive stacks provide linear peak pressures across the range of 2-4.5 peak psi. Representative pressure versus time (milliseconds) profiles averaged from 3-6 shocktube blasts (red) as compared to the idealized Friedlander curves (blue) for (B) 1 sheet, (C) 2 sheets, (D) 3 sheets, and (E) 4 sheets. <a href="https://www.jove.com/files/ftp_upload/66592/66592fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66592/66592fig03.jpg" /> 
Figure 3: Intersubject Interval. Set up and execution of a single blast requires on average 9.8 &#177; 1.9 min (mean &#177; standard error of the mean (sem)). Additional blast exposures require an additional 1.7 &#177; 0.4 min per event (mean &#177; sem). Dots represent results from individual animals. <a href="https://www.jove.com/files/ftp_upload/66592/66592fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66592/66592fig04.jpg" /> 
Figure 4: Daily righting times during 3 weeks of highly repetitive LLB exposures. The graph represents the sham-normalized righting times over 3 weeks of LLB exposure. LLB mice were subject to 6 daily blast exposures for a total of 90 total LLB exposures occurring over 15 days. Mean overpressure characteristics were (&#177; sem) 3.05 &#177; 0.07 peak psi, 0.94 &#177; 0.04 positive phase duration, and 2 &#177; 0.1 psi * msec impulse. p-values reflect results from 2-way ANOVA. Abbreviation: LLB = Low-level blast. <a href="https://www.jove.com/files/ftp_upload/66592/66592fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/66592/66592fig05.jpg" /> 
Figure 5: Effects of the laboratory shocktube LLB model on animal attrition following highly repetitive LLB exposures. Attrition rates for sham (N = 24) and LLB mice (N = 32) from the first LLB exposure (day 1) through all study exposures (ending day 19) and following a 6-month recovery period (day 199). There was no significant difference between the attrition rates of sham and LLB groups over the observed period. LLB mice experienced an average of 62 exposures at an average of 4.78 &#177; 0.01 peak psi and 3.16 &#177; 0.023 psi&#8729;ms impulse. Exposures were administered to mice 5 days per week (i.e., Monday-Friday) for 3 consecutive weeks to model recently reported SOF overpressure exposures during routine breaching training45. Abbreviation: LLB = Low-level blast; SOF = Special Operations Forces. <a href="https://www.jove.com/files/ftp_upload/66592/66592fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Author Spotlight: Deciphering the Long-Term Effects of Low-Level Blast Exposures in Mice

Presented here are methods for producing repeated low-intensity blast exposures using mice.

Modeling Highly Repetitive Low-level Blast Exposure in Mice

Exposure to explosive blasts is a significant risk factor for brain trauma among exposed persons. Although the effects of large blasts on the brain are ...

Our research focuses on the long-term effects of repeated low level blast exposure in mice. Our goal is to determine the long-term effects of this exposure during military training For animal research, the effects of low level blast are subtle, and research teams will often use techniques like electron microscopy to detect differences between blast and sham mice. Our work suggests increased blast repetition makes these differences easier to detect using more accessible microscopy techniques.There's currently very limited literature on both low level blast exposures, which are also called low intensity blasts, and the models that use these exposures over multiple days. We designed our protocol to include multiple low level blasts over many days to address these gaps.

modeling-highly-repetitive-low-level-blast-exposure-in-mice

Research

JoVE Journal

Neuroscience

472 Views.  United States Army Special Operations Command. Our research focuses on the long-term effects of repeated low level blast exposure in mice. Our goal is to determine the long-term effects of this exposure during military training For animal research, the effects of low level blast are subtle, and research teams will often use techniques like electron microscopy to detect differences between blast and sham mice. Our work suggests increased blast repetition makes these differences easier to detect using more accessible microscopy techniques.

Video: Author Spotlight: Deciphering the Long-Term Effects of Low-Level Blast Exposures in Mice

Modeling Astrocytoma Pathogenesis In Vitro and In Vivo Using Cortical Astrocytes or Neural Stem Cells from Conditional, Genetically Engineered Mice

Current astrocytoma models are limited in their ability to define the roles of oncogenic mutations in specific brain cell types during disease pathogenesis and their utility for preclinical drug development. In order to design a better model system for these applications, phenotypically wild-type cortical astrocytes and neural stem cells (NSC) from conditional, genetically engineered mice (GEM) that harbor various combinations of floxed oncogenic alleles were harvested and grown in culture. Genetic recombination was induced in vitro using adenoviral Cre-mediated recombination, resulting in expression of mutated oncogenes and deletion of tumor suppressor genes. The phenotypic consequences of these mutations were defined by measuring proliferation, transformation, and drug response in vitro. Orthotopic allograft models, whereby transformed cells are stereotactically injected into the brains of immune-competent, syngeneic littermates, were developed to define the role of oncogenic mutations and cell type on tumorigenesis in vivo. Unlike most established human glioblastoma cell line xenografts, injection of transformed GEM-derived cortical astrocytes into the brains of immune-competent littermates produced astrocytomas, including the most aggressive subtype, glioblastoma, that recapitulated the histopathological hallmarks of human astrocytomas, including diffuse invasion of normal brain parenchyma. Bioluminescence imaging of orthotopic allografts from transformed astrocytes engineered to express luciferase was utilized to monitor in vivo tumor growth over time. Thus, astrocytoma models using astrocytes and NSC harvested from GEM with conditional oncogenic alleles provide an integrated system to study the genetics and cell biology of astrocytoma pathogenesis in vitro and in vivo and may be useful in preclinical drug development for these devastating diseases.

Modeling Astrocytoma Pathogenesis In Vitro and In Vivo Using Cortical Astrocytes or Neural Stem Cells from Conditional, Genetically Engineered Mice

Phenotypically wild-type astrocytes and neural stem cells harvested from mice engineered with floxed, conditional oncogenic alleles and transformed via viral Cre-mediated recombination can be used to model astrocytoma pathogenesis in vitro and in vivo by orthotopic injection of transformed cells into brains of syngeneic, immune-competent littermates.

Current astrocytoma models are limited in their ability to define the roles of oncogenic mutations in specific brain cell types during disease ...

A Simple and Inexpensive Method for Determining Cold Sensitivity and Adaptation in Mice

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life. The cold plantar assay allows the objective and inexpensive assessment of cold sensitivity in mice, and can quantify both analgesia and hypersensitivity. Mice are acclimated on a glass plate, and a compressed dry ice pellet is held against the glass surface underneath the hindpaw. The latency to withdrawal from the cooling glass is used as a measure of cold sensitivity.
Cold sensation is also important for survival in regions with seasonal temperature shifts, and in order to maintain sensitivity animals must be able to adjust their thermal response thresholds to match the ambient temperature. The Cold Plantar Assay (CPA) also allows the study of adaptation to changes in ambient temperature by testing the cold sensitivity of mice at temperatures ranging from 30 &#176;C to 5 &#176;C. Mice are acclimated as described above, but the glass plate is cooled to the desired starting temperature using aluminum boxes (or aluminum foil packets) filled with hot water, wet ice, or dry ice. The temperature of the plate is measured at the center using a filament T-type thermocouple probe. Once the plate has reached the desired starting temperature, the animals are tested as described above.
This assay allows testing of mice at temperatures ranging from innocuous to noxious. The CPA yields unambiguous and consistent behavioral responses in uninjured mice and can be used to quantify both hypersensitivity and analgesia. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

The Cold Plantar Assay (CPA) measures cold responsiveness between 30 &#176;C and 5 &#176;C, and can also measure cold adaptation. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life.  The cold ...

Modeling Neuronal Death and Degeneration in Mouse Primary Cerebellar Granule Neurons

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their post-natal development, abundance, and accessibility, CGNs are an ideal model to study neuronal processes, including neuronal development, neuronal migration, and physiological neuronal activity stimulation. In addition, CGN cultures provide an excellent model for studying different modes of cell death including excitotoxicity and apoptosis. Within a week in culture, CGNs express N-methyl-D-aspartate (NMDA) receptors, a specific ionotropic glutamate receptor with many critical functions in neuronal health and disease. The addition of low concentrations of NMDA in conjunction with membrane depolarization to rodent primary CGN cultures has been used to model physiological neuronal activity stimulation while the addition of high concentrations of NMDA can be employed to model excitotoxic neuronal injury. Here, a method of isolation and culturing of CGNs from 6 day old pups as well as genetic manipulation of CGNs by adenoviruses and lentiviruses are described. We also present optimized protocols on how to stimulate NMDA-induced excitotoxicity, low-potassium-induced apoptosis, oxidative stress and DNA damage following transduction of these neurons.

This protocol describes a simple method for isolating and culturing primary mouse cerebral granule neurons (CGNs) from 6-7 day old pups, efficient transduction of CGNs for loss and gain of function studies, and modelling NMDA-induced neuronal excitotoxicity, low-potassium-induced cell death, DNA-damage, and oxidative stress using the same culture model.

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their ...

A Novel In Vitro Model of Blast Traumatic Brain Injury

Traumatic brain injury is a leading cause of death and disability in military and civilian populations. Blast traumatic brain injury results from the detonation of explosive devices, however, the mechanisms that underlie the brain damage resulting from blast overpressure exposure are not entirely understood and are believed to be unique to this type of brain injury. Preclinical models are crucial tools that contribute to better understand blast-induced brain injury. A novel in vitro blast TBI model was developed using an open-ended shock tube to simulate real-life open-field blast waves modelled by the Friedlander waveform. C57BL/6N mouse organotypic hippocampal slice cultures were exposed to single shock waves and the development of injury was characterized up to 72 h using propidium iodide, a well-established fluorescent marker of cell damage that only penetrates cells with compromised cellular membranes. Propidium iodide fluorescence was significantly higher in the slices exposed to a blast wave when compared with sham slices throughout the duration of the protocol. The brain tissue injury is very reproducible and proportional to the peak overpressure of the shock wave applied.

A Novel In Vitro Model of Blast Traumatic Brain Injury

This paper describes a novel model of primary blast traumatic brain injury. A compressed-air driven shock tube is used to expose in vitro mouse hippocampal slice cultures to a single shock wave. This is a simple and rapid protocol generating a reproducible brain tissue injury with a high throughput.

Traumatic brain injury is a leading cause of death and disability in military and civilian populations. Blast traumatic brain injury results from the ...

Quantitative Measurement of Intrathecally Synthesized Proteins in Mice

Cerebrospinal fluid (CSF), a fluid found in the brain and the spinal cord, is of great importance to both basic and clinical science. The analysis of the CSF protein composition delivers crucial information in basic neuroscience research as well as neurological diseases. One caveat is that proteins measured in CSF may derive from both intrathecal synthesis and transudation from serum, and protein analysis of CSF can only determine the sum of these two components. To discriminate between protein transudation from the blood and intrathecally produced proteins in animal models as well as in humans, CSF protein profiling measurements using conventional protein analysis tools must include the calculation of the albumin CSF/serum quotient (Qalbumin), a marker of the integrity of the blood-brain interface (BBI), and the protein index (Qprotein/Qalbumin), an estimate of intrathecal protein synthesis. This protocol illustrates the entire procedure, from CSF and blood collection to quotients and indices calculations, for the quantitative measurement of intrathecal protein synthesis and BBI impairment in mouse models of neurological disorders.

Elevated spinal fluid protein levels can either be the result of diffusion of plasma protein across an altered blood-brain barrier or intrathecal synthesis. An optimized testing protocol is presented in this article that helps to discriminate both cases and provides quantitative measurements of intrathecally synthesized proteins.

Cerebrospinal fluid (CSF), a fluid found in the brain and the spinal cord, is of great importance to both basic and clinical science. The analysis of the ...

Modeling Stroke in Mice: Focal Cortical Lesions by Photothrombosis

Stroke is a leading cause of death and acquired adult disability in developed countries. Despite extensive investigation for novel therapeutic strategies, there remain limited therapeutic options for stroke patients. Therefore, more research is needed for pathophysiological pathways such as post-stroke inflammation, angiogenesis, neuronal plasticity, and regeneration. Given the inability of in vitro models to reproduce the complexity of the brain, experimental stroke models are essential for the analysis and subsequent evaluation of novel drug targets for these mechanisms. In addition, detailed standardized models for all procedures are urgently needed to overcome the so-called replication crisis. As an effort within the ImmunoStroke research consortium, a standardized photothrombotic mouse model using an intraperitoneal injection of Rose Bengal and the illumination of the intact skull with a 561 nm laser is described. This model allows the performance of stroke in mice with allocation to any cortical region of the brain without invasive surgery; thus, enabling the study of stroke in various areas of the brain. In this video, the surgical methods of stroke induction in the photothrombotic model along with&#160;histological analysis are demonstrated.

Described here is the photothrombotic stroke model, where a stroke is produced through the intact skull by inducing permanent microvascular occlusion using laser illumination after administration of a photosensitive dye.

Stroke is a leading cause of death and acquired adult disability in developed countries. Despite extensive investigation for novel therapeutic strategies, ...

Modeling Stroke in Mice: Transient Middle Cerebral Artery Occlusion via the External Carotid Artery

Stroke is the third most common cause of mortality and the leading cause of acquired adult disability in developed countries. To date, therapeutic options are limited to a small proportion of stroke patients within the first hours after stroke. Novel therapeutic strategies are being extensively investigated, especially to prolong the therapeutic time window. These current investigations include the study of important pathophysiological pathways after stroke, such as post-stroke inflammation, angiogenesis, neuronal plasticity, and regeneration. Over the last decade, there has been increasing concern about the poor reproducibility of experimental results and scientific findings among independent research groups. To overcome the so-called &#8220;replication crisis&#8221;, detailed standardized models for all procedures are urgently needed. As an effort within the &#8220;ImmunoStroke&#8221; research consortium (https://immunostroke.de/), a standardized mouse model of transient middle cerebral artery occlusion (MCAo) is proposed. This model allows the complete restoration of the blood flow upon removal of the filament, simulating the therapeutic or spontaneous clot lysis that occurs in a large proportion of human strokes. The surgical procedure of this &#8220;filament&#8221; stroke model and tools for its functional analysis are demonstrated in the accompanying video.

Different models of middle cerebral artery occlusion (MCAo) are used in experimental stroke research. Here, an experimental stroke model of transient MCAo via the external carotid artery (ECA) is described, which aims to mimic human stroke, in which the cerebrovascular thrombus is removed due to spontaneous clot lysis or therapy.

Stroke is the third most common cause of mortality and the leading cause of acquired adult disability in developed countries. To date, therapeutic options ...

A Pilot Study on the Repetitive Transcranial Magnetic Stimulation of A&#946; and Tau Levels in Rhesus Monkey Cerebrospinal Fluid

Previous studies have demonstrated that a non-invasive light-flickering regime and auditory tone stimulation could affect A&#946; and tau metabolism in the brain. As a non-invasive technique, repetitive transcranial magnetic stimulation (rTMS) has been applied for the treatment of neurodegenerative disorders. This study explored the effects of rTMS on A&#946; and tau levels in rhesus monkey cerebrospinal fluid (CSF). This is a single-blind, self-controlled study. Three different frequencies (low frequency, 1 Hz; high frequencies, 20 Hz and 40 Hz) of rTMS were used to stimulate the bilateral-dorsolateral prefrontal cortex (DLPFC) of the rhesus monkey. A catheterization method was used to collect CSF. All samples were subjected to liquid chip detection to analyze CSF biomarkers (A&#946;42, A&#946;42/A&#946;40, tTau, pTau). CSF biomarker levels changed with time after stimulation by rTMS. After stimulation, the A&#946;42 level in CSF showed an upward trend at all frequencies (1 Hz, 20 Hz, and 40 Hz), with more significant differences for the high-frequencies (p &lt; 0.05) than for the low frequency. 
After high-frequency rTMS, the total Tau (tTau) level of CSF immediately increased at the post-rTMS timepoint (p &lt; 0.05) and gradually decreased by 24 h. Moreover, the results showed that the level of phosphorylated Tau (pTau) increased immediately after 40 Hz rTMS (p &lt; 0.05). The ratio of A&#946;42/A&#946;40 showed an upward trend at 1 Hz and 20 Hz (p &lt; 0.05). There was no significant difference in the tau levels with low-frequency (1 Hz) stimulation. Thus, high-frequencies (20 Hz and 40 Hz) of rTMS may have positive effects on A&#946; and tau levels in rhesus monkey CSF, while low-frequency (1 Hz) rTMS can only affect A&#946; levels.

Here, we describe the procedure for a pilot study to explore the effect of repetitive transcranial magnetic stimulation with different frequencies (1 Hz/20 Hz/40 Hz) on A&#946; and tau metabolism in rhesus monkey cerebrospinal fluid.

Previous studies have demonstrated that a non-invasive light-flickering regime and auditory tone stimulation could affect A&#946; and tau metabolism in ...

Modeling Neonatal Intraventricular Hemorrhage Through Intraventricular Injection of Hemoglobin

Neonatal intraventricular hemorrhage (IVH) is a common consequence of premature birth and leads to brain injury, posthemorrhagic hydrocephalus (PHH), and lifelong neurological deficits. While PHH can be treated by temporary and permanent cerebrospinal fluid (CSF) diversion procedures (ventricular reservoir and ventriculoperitoneal shunt, respectively), there are no pharmacological strategies to prevent or treat IVH-induced brain injury and hydrocephalus. Animal models are needed to better understand the pathophysiology of IVH and test pharmacological treatments. While there are existing models of neonatal IVH, those that reliably result in hydrocephalus are often limited by the necessity for large-volume injections, which may complicate modeling of the pathology or introduce variability in the clinical phenotype observed.
Recent clinical studies have implicated hemoglobin and ferritin in causing ventricular enlargement after IVH. Here, we develop a straightforward animal model that mimics the clinical phenotype of PHH utilizing small-volume intraventricular injections of the blood breakdown product hemoglobin. In addition to reliably inducing ventricular enlargement and hydrocephalus, this model results in white matter injury, inflammation, and immune cell infiltration in periventricular and white matter regions. This paper describes this clinically relevant, simple method for modeling IVH-PHH in neonatal rats using intraventricular injection and presents methods for quantifying ventricle size post injection.

We present a model of neonatal intraventricular hemorrhage using rat pups that mimics the pathology seen in humans.

Neonatal intraventricular hemorrhage (IVH) is a common consequence of premature birth and leads to brain injury, posthemorrhagic hydrocephalus (PHH), and ...

Modeling Human Cerebellar Development In Vitro in 2D Structure

The precise and timely development of the cerebellum is crucial not only for accurate motor coordination and balance but also for cognition. In addition, disruption in cerebellar development has been implicated in many neurodevelopmental disorders, including autism, attention deficit-hyperactivity disorder (ADHD), and schizophrenia. Investigations of cerebellar development in humans have previously only been possible through post-mortem studies or neuroimaging, yet these methods are not sufficient for understanding the molecular and cellular changes occurring in vivo during early development, which is when many neurodevelopmental disorders originate. The emergence of techniques to generate human-induced pluripotent stem cells (iPSCs) from somatic cells and the ability to further re-differentiate iPSCs into neurons have paved the way for in vitro modeling of early brain development. The present study provides simplified steps toward generating cerebellar cells for applications that require a 2-dimensional (2D) monolayer structure. Cerebellar cells representing early developmental stages are derived from human iPSCs via the following steps: first, embryoid bodies are made in 3-dimensional (3D) culture, then they are treated with FGF2 and insulin to promote cerebellar fate specification, and finally, they are terminally differentiated as a monolayer on poly-l-ornithine (PLO)/laminin-coated substrates. At 35 days of differentiation, iPSC-derived cerebellar cell cultures express cerebellar markers including ATOH1, PTF1&#945;, PAX6, and KIRREL2, suggesting that this protocol generates glutamatergic and GABAergic cerebellar neuronal precursors, as well as Purkinje cell progenitors. Moreover, the differentiated cells show distinct neuronal morphology and are positive for immunofluorescence markers of neuronal identity such as TUBB3. These cells express axonal guidance molecules, including semaphorin-4C, plexin-B2, and neuropilin-1, and could serve as a model for investigating the molecular mechanisms of neurite outgrowth and synaptic connectivity. This method generates human cerebellar neurons useful for downstream applications, including gene expression, physiological, and morphological studies requiring 2D monolayer formats.

Modeling Human Cerebellar Development In Vitro in 2D Structure

The present protocol explains the generation of a 2D monolayer of cerebellar cells from induced pluripotent stem cells for investigating the early stages of cerebellar development.

The precise and timely development of the cerebellum is crucial not only for accurate motor coordination and balance but also for cognition. In addition, ...