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Aug 30, 2023

CDk5 は回転力を媒介します

Rapporti scientifici Volume 13,

Scientific Reports volume 13, Article number: 3394 (2023) Cite this article

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Millions of traumatic brain injuries (TBIs) occur annually. TBIs commonly result from falls, traffic accidents, and sports-related injuries, all of which involve rotational acceleration/deceleration of the brain. During these injuries, the brain endures a multitude of primary insults including compression of brain tissue, damaged vasculature, and diffuse axonal injury. All of these deleterious effects can contribute to secondary brain ischemia, cellular death, and neuroinflammation that progress for weeks, months, and lifetime after injury. While the linear effects of head trauma have been extensively modeled, less is known about how rotational injuries mediate neuronal damage following injury. Here, we developed a new model of repetitive rotational head trauma in rodents and demonstrated acute and prolonged pathological, behavioral, and electrophysiological effects of rotational TBI (rTBI). We identify aberrant Cyclin-dependent kinase 5 (Cdk5) activity as a principal mediator of rTBI. We utilized Cdk5-enriched phosphoproteomics to uncover potential downstream mediators of rTBI and show pharmacological inhibition of Cdk5 reduces the cognitive and pathological consequences of injury. These studies contribute meaningfully to our understanding of the mechanisms of rTBI and how they may be effectively treated.

Traumatic brain injuries occur in epidemic proportions with over 1.5 million injuries occurring annually in the United States alone1. During a TBI, the brain undergoes both linear and rotational forces, compressing tissue and damaging axonal connections. While the linear effects of injury primarily result in focal brain damage, rotational acceleration of the brain produces shearing forces resulting in both focal and diffuse neuropathology. Diffuse axonal shearing as the result of rotational TBI (rTBI) causes persistent neuropathological consequences of brain trauma2,3,4. At the time of injury, rapid brain acceleration/deceleration creates a disruption of brain tissue resulting in contusion, blood vessel damage, hemorrhage, and axonal shearing5. TBI results in acute dysfunctions in cognition including loss of consciousness, headaches, and vison problems6. Secondary damage evolves over the following days, months, and lifetime of survivors leading to impairments in memory and motor function, anxiodepressive disorders, and neurodegeneration7,8,9,10.

At cellular resolution, rTBI induces axonal shearing resulting in massive neuronal depolarization and ionic influx11,12,13,14,15. In response, activation of voltage gated Ca2+ channels induce excitotoxic release of glutamate. Following excitotoxicity, cerebral edema, oxidative stress, and cellular death all contribute to the acute phase of injury1,16,17. After initial trauma, a delayed spreading process of injury occurs. The injured brain exhibits increased sensitivity to secondary ischemic insult and persistent excitotoxicity18. Secondary damage advances over weeks, months, and years after injury, impairing neurological function and impeding recovery19. Chronic effects from TBI include increased risk of developing Alzheimer's disease (AD), increased risk of developing Parkinson's disease (PD), and neurodegeneration in the form of chronic traumatic encephalopathy (CTE)20,21,22.

The disruption of intracellular signaling cascades serves as a convergence point of ischemia, inflammation, and excitotoxicity where the activation of downstream effectors induces cellular demise. One such excitotoxic pathway is the dysregulation of the protein kinase cyclin dependent kinase 5 (Cdk5) via the calpain family of Ca2+-activated neutral proteases23. Following cellular damage, activated calpain protease cleaves the coactivator of Cdk5, p35, into a truncated aberrant coactivator, p2524. Conversion of Cdk5/p35 to Cdk5/p25 by calpain confers neurotoxic activity upon the kinase, resulting in neuronal injury and death25. Aberrant Cdk5/p25 contributes to virtually all neurodegenerative diseases and is key to the general processes by which neurotoxicity occurs25,26,27. Conditional knockout of Cdk5 confers neuroprotection from cortical impact, ischemic stroke, and mouse models of AD23,28,29. Therefore, we hypothesized that aberrant Cdk5/p25 activity may mediate the neuropathological and neurocognitive effects of repeated and rotational brain injuries. To test this, we designed and characterized a novel model of rTBI and assessed the neuropathological effects of repetitive injury in the acute and subacute phases, including aberrant Cdk5/p25 activation. We assessed a new systemic Cdk5 inhibitor (25–106) as a potential treatment for rTBI. Finally, we conducted Cdk5-enriched phosphoproteomics to identify novel downstream effectors of brain injury. Together, these studies provide a better understanding of the mechanisms mediating injury by rTBI and point to a potentially effective therapeutic approach.

To assess the negative consequences of rotational head injury, we developed a novel repetitive rTBI model with the ability to impart clinically relevant angular accelerations (Fig. 1A). The model consists of a pneumatic-chain driven pendulum arm attached to an animal carrier. This pendulum rotates about a horizontal shaft. The rodent is placed in a restraint inserted onto a fixed mount at the end of the pendulum (Fig. 1A, B) and attached to the freely rotating helmet assembly bolted to the restraint mount (Fig. 1C), allowing pure coronal plane head rotation. A ventral strike plate (Fig. 1D,) transfers impact momentum between the harness and impact fixture, imparting rotational force on the subject's head. The pendulum is rapidly propelled forward using a compressed N2-driven 2-way solenoid valve (Fig. 1E, F) that subsequently drives an air motor inducing rapid chain rotation. Accelerometry is derived from a helmet mounted inertial measurement unit (IMU) model 633, a 6-degrees of freedom transducer incorporating accelerometers and gyroscopes comprised of micro-machined silicon sensors (Fig. 1G, H). IMU data is captured in LabVIEW and converted to acceleration measured in gravitational constants (g values), which are converted to angular acceleration (rad/sec2).

Novel rodent model of rotational traumatic brain injury (rTBI). (A) Rotational head injury model, components, and complete design. (B) Animal restraint. (C) Helmet and subject constraint mount. (D) Ventral strike plate. E. Pneumatic-chain drive system. (F) Pneumatic pressure system. (G) Side view of helmet assembly and restraint with artificial rat. Note model 633 IMU mounted on helmet strike plate below rat head with front view showing strike plate and impact bumper. (H) Model 633 6DOF inertia measurement unit. Oscillating variable capacitance gyro sensor combs (right) and half cover removed view of IMU showing acceleration sensor, mass, supports, and circuitry. (I) Temporally sequential high- speed camera recording frames used to derive acceleration rates. (J) Sample recording from IMU. (K) Video telemetry calculated rate vs. time. (L) Overlay of helmet IMU and video telemetry angular speeds (o/S) as a function of time. (M) Labview derived rotational accelerations (Krad/S2) of 5 trial runs (left). Peak rotational accelerations for each run with standard error (right).

To validate helmet sensor accelerometry, we utilized high speed video telemetry to determine angular speed of the helmet and compared these values to those derived from the helmet sensor (Fig. 1I–L). Video telemetry and helmet sensor showed highly overlapping determinates for angular speed. Typical machine runs induce average helmet peak rotational acceleration outputs averaging 482Krad/s2 with limited variability across 5 sampling runs of 14.15 Krad/s2 (Fig. 1M). These angular accelerations are consistent with human to animal scaling laws for mild brain injuries30,31, suggesting rTBI studies using this model in rodents may readily translate to human conditions.

The consequences of rTBI include acute and prolonged pathological alterations including damaged vasculature, diffuse axonal injury, inflammation, cellular demise, and neurodegeneration5. To investigate the effects of rTBI, we first conducted PET/CT neuropathology studies. For these studies, we made use of a diagnostic molecular probe in current clinical use for the detection of the neuroinflammatory marker, translocator protein (TSPO). This probe, [18F]DPA-714 detects activated microglia resulting from neuroinflammation in neurogenerative and neuroinflammatory conditions32,33,34. TSPO imaging revealed differential uptake diffusely throughout the brains of both control and rTBI rats 7 days post-injury (Fig. 2A). Interestingly, standard uptake values (SUV) were significantly increased in several brain regions including amygdala, hippocampus CA1 layer, and the basolateral amygdaloid nucleus of rTBI rats 7 days post-injury in comparison to controls (Fig. 2B–D). Effects were also observed diffusely throughout whole cortex with cortical areas including perirhinal and primary somatosensory cortex showing higher labeling following rTBI (Supplemental Fig. 1A–C). The differential labeling was region-specific as no effect was detectible when signal throughout whole brain was quantitated (Supplemental Fig. 1D). Also, regions such as hippocampus layer CA3 and cerebellum exhibited no change in response to impact (Supplemental Fig. 1E, F). Thus, rTBI caused brain region-specific alterations in this biomarker of neuroinflammation, consistent with effects observed in humans32.

Neuropathological effects of rTBI. (A) Representative PET/CT images for TSPO uptake in control (left) and rTBI (right) rats (n = 6 per group). (B) Quantitative Standard uptake values (SUV) for TSPO radioligand in amygdala (Time: F(20,200) = 110.6, p < 0.0001; Treatment: (1, 10) = 7.771, p = 0.0192; Interaction: (20, 200) = 2.321, p = 0.0017) two-way-RM ANOVA. (C) CA1 (Time: F(5,50) = 216.4, p < 0.0001; Treatment: F(1,10) = 1.711, p = 0.2202; Interaction: F(5,50) = 2.614, p = 0.0355) two-way-RM ANOVA. (D) Basolateral amygdaloid nucleus (Time: F(10,100) = 4 9.67, p < 0.0001; Treatment: F(1,10) = 13.97, p = 0.0039; Interaction: F(10,100) = 2.605, p = 0.0075). (E) Histological staining and quantitation for IBA1+ microglia within CA1, p = 0.0083 Student's t-test. (n = 4–5 per group) Scale bars = 50 µm. (F) Histological staining and quantitation for GFAP+ astrocytes within CA1, p = 0.0317 Mann Whitney test (p = 0.0300, Shapiro–Wilk) (n = 4–5 per group) Scale bars = 50 µm. (G) Histological staining for AT8+ (Top, left) and condensed staining around vessels (Top, right) Scale bars = 100 µm. Histological staining and quantification of AT8 density in CA1 at 6 months post injury (bottom) p = 0.0210 Student's t-test (n = 5 per group) Scale bars = 20 µm. (H) Histological staining for SMI-31 at 48 h and 14 days post injury. (n = 4–5 per group) (48 h Con-TBI, p = 0.0409 Student's t-test). All data are means ± SEM, *p < 0.05, ** < 0.01, ** < 0.01, **** < 0.001.

In addition to in vivo imaging, immunostaining was used to assess neuroinflammatory effects of rTBI. For these experiments, layers of the hippocampal formation were examined, as rTBI effects within this brain region in humans have been linked to consequent memory impairments35,36,37. Indeed, broad increases in microgliosis were detected throughout the CA1 subfield of the hippocampus (Fig. 2E) 48 h post-injury. Furthermore, rats subjected to rTBI showed increased astrogliosis throughout the CA1 (Fig. 2F). Chronic consequences of brain injury include tauopathy occurring in CTE38. To assess chronic neuropathological changes following rTBI, rats 6 months post injury were assessed for increased phospho-Tau (AT8) immunoreactivity. Remarkably, rotational injury induced an increase in AT8 immunoreactivity throughout the brain at 6 months post-injury particularly in areas surrounding vasculature (Fig. 2G, Top). We also observed increased AT8 staining, with diffuse staining throughout the neuropil in regions of the CA1 in rTBI rats as compared to age-matched controls (Fig. 2G, Bottom). Another pathological hallmark of TBI is axonal injury39. To assess any structural damage to axons of the hippocampus, we stained the CA1 subregion for phosphorylated neurofilaments (SMI-31), a marker of axonal injury40. We observed an increased in SMI-31 48 h after injury, but not 14 days after rTBI (Fig. 2H). Together, these in vivo imaging and histological studies demonstrate rotational head injury induces neuroinflammation and axonal damage in acute and sub-acute phases of injury. These effects underlie prolonged changes in AT8 immunoreactivity.

Persistent memory impairment is a common outcome following TBI8. Patients report symptoms including both anterograde and retrograde amnesia, as well as cognitive deficits in attention, processing speed, and executive functioning41. These deficits often correlate with structural damage to temporal brain areas35 and damage within the hippocampal formation following TBI can contribute to long-lasting deficits in memory42. Therefore, we investigated the functional and physiological consequences of rTBI on hippocampal-dependent memory and plasticity in our model. To assess effects on contextual and cued fear learning and memory, rats (7 days post-injury) were placed in a novel context, analyzed for baseline freezing responses, and subsequently exposed to fear conditioning cue/foot shock pairings for assessment of learning and memory function (Fig. 3A). Rats subjected to rTBI displayed no alteration in baseline freezing rates. However, rTBI rats showed a marked decrease in freezing behavior in response to re-exposure to the adverse context. Additionally, rTBI rats displayed a reduction in cue-induced freezing rates in a novel context, suggesting that both hippocampal- and amygdala-mediated components of this learned behavior were impaired. This reduction in freezing responses was not confounded by injury-induced damage to nociceptive circuitry, as shock stimulus pain thresholds for animal flinching, jumping, and vocalizing pain were not altered by rTBI (Fig. 3B). Thus, rTBI caused learning and memory deficits that persisting into the sub-acute phase of injury.

Behavioral and neurophysiological consequences of rTBI. (A) Fear conditioning freezing rates for baseline, Contextual, p = 0.0001 Student's t-test, and Cued, p = 0.0018 Student's t-test fear learning (n = 8–11 per group). (B) Shock sensitivity thresholds to flinch, jump, or vocalize pain (n = 8–11 per group). (C) Input–Output curve of CA3-CA1 fEPSP recordings (Inset: individual traces for each of the stimulus intensities) (n = 5–6 per group). (D) Paired Pulse Ratio (PPR) across inter-stimulus interval (n = 5–6 per group). (E) Assessment of the effect of rTBI on hippocampal plasticity after HFS (n = 5–6 per group). (PTP outlined grey 0–2 min, LTP outlined grey 45–55 min). (F) Summary of PTP and LTP fEPSP slopes, (n = 5–6 per group) p = 0.7662, PTP; p = 0.0017, LTP Student's t-test. All data are means ± SEM, *p < 0.05, ** < 0.01, *** < 0.001.

The observed mnemonic impairments in response to rTBI suggest that hippocampal circuitry function may be damaged. To investigate this possibility, field excitatory post synaptic potential (fEPSP) recordings were taken from hippocampal CA3-CA1 circuitry (7 days post-injury) (Fig. 3C–F). Synaptic excitability, as assessed via stimulus-to-fEPSP amplitude ratios (I/V curves), was unaffected by rTBI (Fig. 3C). The paired-pulse ratio paradigm was used to assess changes on neurotransmitter release. Similarly, this metric of short-term plasticity remained unaltered in rTBI rats (Fig. 3D). Additionally, high frequency stimulation (HFS) induced a 171.4 ± 36% post-tetanic potentiation (PTP) in control rats compared to baseline (Fig. 3E, F). This effect was similar (183.4 ± 18%) in rTBI brains. HFS also induced robust (300 ± 47%) long-term potentiation (LTP) of fEPSP slope compared to baseline 45 min post-stimulus at Schaffer collateral-CA1 synapses of control mice. (Fig. 3E, F). In contrast, tetanic stimulation caused a markedly smaller effect (106 ± 8%) in rTBI rats, almost completely ablating the ability to induce LTP. Thus, rTBI caused a notable impairment in hippocampal long-term synaptic plasticity as a likely mechanism underlying the deficit it induced in learning and memory.

TBI may impart long-lasting impairments through alterations in protein expression that lead to cell death or inflammatory pathways ultimately leading to neuronal demise. To ascertain the effects of rTBI on protein levels as a net measure of the balance between expression and degradation, we conducted global proteomics to identify mechanisms of rTBI associated with the acute inflammatory state of the hippocampus we observed at 48 h post-injury (Fig. 4A). We noted 234 proteins demonstrated increased expression (FC ≥ 1.3) and 250 proteins with decreased expression (FC ≤ − 1.3) (Fig. 4B) in rTBI hippocampus relative to controls. The altered expression observed in the proteomic analysis within rTBI lysates was independently validated for two specific proteins (Supplemental Fig. 2A, B). Proteomic analysis revealed animals subjected to rTBI displayed a 1.6 FC of Legumain (LGMN) protease following injury (Fig. 4B). Immunoblot analysis of hippocampal lysates confirmed LGMN is significantly upregulated in expression following injury (Supplemental Fig. 2A). Additionally, Epoxide hydroxylase 2 (EPHX2) displayed a FC increase of 3.9 in rTBI rats compared to control rats. Similarly, immunoblot analysis displayed significant increases of EPHX2 in hippocampal lysates following injury (Supplemental Fig. 2B).

Proteomic alterations following rTBI. (A) Experimental design schematic for tissue isolation, extraction, labeling, and peptide detection for discovery proteomics. (B) Volcano plot of differentially expressed total proteins between rTBI and CON (control) rats (n = 5 pooled brains in duplicate per group, increased sites in green decreased sites in red). (C) (Top) Ingenuity Pathway Analysis of upregulated proteins following rTBI. (Bottom) Ingenuity Pathway Analysis of downregulated proteins following rTBI. (D) Cell-type expression profile in brain of differentially expressed proteins assessed from available dataset47. Abbreviations: (ABC) Arachnoid barrier cells, (mNEUR) Mature neurons, (ARP) Astrocyte-restricted precursors, (NendC) Neuroendocrine cells, (ASC) Astrocytes, (NEUT) Neutrophils, (CPC) Choroid plexus epithelial cells, (NRP) Neuronal-restricted precursors, (DC) Dendritic cells, (NSC) Neural stem cells, (EC) Endothelial cells, (OEG) Olfactory ensheathing glia (EPC) Ependymocytes, (OLG) Oligodendrocytes, (Hb_VC) Hemoglobin-expressing vascular cells, (OPC)Oligodendrocyte precursor cells, (HypEPC) Hypendymal cells, (PC) Pericytes (ImmN) Immature neurons, (TNC) Tanycytes, (MAC) Macrophages, (VLMC) Vascular and leptomeningeal cells, (MG) Microglia, (VSMC) Vascular smooth muscle cells, (MNC) Monocytes.

To identify the intracellular pathways altered via rTBI we conducted ingenuity pathways analysis of the top upregulated and downregulated proteins following rTBI (Fig. 4C). Interestingly, rTBI induced upregulation of known cell death dependent pathways linked to brain injury such as ferroptosis and phagosome formation43,44. We observed bidirectional alterations in homeostatic signaling pathways such as Protein Kinase A (PKA) and oxytocin signaling pathways, both of which have been observed to mediate injury in stroke and TBI45,46.

Downregulated pathways after injury included proteins associated with inflammatory response elements such as acute phase response signaling were downregulated in the hippocampus following rTBI. Notably, neurotropic signaling such as those involving neuregulin and wound healing pathways were also downregulated following rTBI. Utilizing an open-source single-cell gene expression atlas of the mouse brain, we determined the cell-type expression profiles of key upregulated proteins following rTBI47 (Fig. 4D). Differentially expressed following rTBI displayed cell-type specific alterations in numerous cell-types within immune cell classes such as microglia (MG), neutrophils (NEUT), dendritic cells (DC), macrophages (MAC), and monocytes (MNC). Proteomic expression was also altered in many vascular cell lineages such as endothelial cells (EC), vascular smooth muscle cells (VSMC), arachnoid barrier cells (ABC), and vascular and leptomeningeal cells (VLMC). Interestingly, we noted of the top differentially expressed proteins assessed, they showed limited or no expression in cell-types of astrocyte lineage such as astrocytes (ASC) and astrocyte restricted precursors (ARP). Likewise, limited proteomic alterations were observed in mature (NEUR_mature), immature (NEUR_immature) and neuroendocrine (NendC) neuronal cell lineages. These data likely indicate global proteomic expression changes are occurring in response to inflammatory immune states and damaged vasculature following rTBI. Altogether, these analyses reveal protein expression changes that may mediate aspects of rTBI and highlight cell types and pathways linked to inflammatory responses which may ultimately contribute to the consequences of injury.

Inflammation due to injury induces excitotoxic states within surrounding neurons, impairing neuronal physiology, neuronal signaling, and ultimately contributing to impaired behavior48. TBI-induced excitotoxicity triggers loss of Ca2+ homeostasis subsequently leading to the activation of calpain proteases and neuronal demise49. A prime target for calpain in neuronal death is the Cdk5 cofactor p35. The activation of calpain and consequent truncation of the Cdk5 coactivator p35 to its aberrant coactivator p2524. The Cdk5/p25 complex has been implicated in various forms of neuronal injury including ischemic stroke, cortical impact, and blast TBI23,28,50. Generation of the aberrantly active Cdk5/p25 complex is neurotoxic51. Cdk5/p25 hyperphosphorylates Tau in models of AD and causes neuronal cell death52. Thus, aberrant Cdk5 invoked by excitotoxicity is a common neuronal injury mechanism shared by many neuropathological conditions28,53.

To better understand the role of aberrant Cdk5 in mediating excitotoxicity in rTBI, we first treated ex vivo brain slices with high concentrations of NMDA (100 μM) and glycine (gly) (50 μM) to induce excitotoxicity. This treatment caused significant generation of p25 (Fig. 5A). While over-expression of p25 is neurotoxic25,52, pharmacological inhibition of Cdk5/p25 activity is neuroprotective in vitro28. Furthermore, conditional knockout of Cdk5 is neuroprotective in vivo23,28. However, effective targeting of Cdk5 as a post-insult therapeutic approach has not been possible due to the lack of brain penetrant Cdk5/p25 inhibitors. Recently, we discovered a brain diffusible Cdk5 inhibitor (25–106) with exponential brain distribution kinetics and 30-fold greater specificity for Cdk5 over other cyclin dependent kinase family members54. We hypothesized that this inhibitor could have rTBI therapeutic potential by blocking aberrant Cdk5/p25 activity. We first verified that this compound acted as a Cdk5 inhibitor in brain tissue by assessing the effect of slice treatment with 25–106 on phosphorylation of the Cdk5 reporter, Thr75 DARPP32 (Fig. 5B). Indeed, this site was strongly attenuated in brain slices by 25–106 treatment. Next, we assessed the neuroprotective capacity of 25–106 under excitotoxic conditions. The neuroprotective effect following Cdk5 inhibition was observed in ex vivo brain slices treated with high concentrations of NMDA/Gly as evidenced by TTC viability staining (Fig. 5C). These data confirm 25–106 as an inhibitor of Cdk5 in brain tissue and suggest it is neuroprotective in an ex vivo model of excitotoxicity.

rTBI and excitotoxicity evoke aberrant Cdk5 activity and Cdk5 inhibition is protective. (A) Quantitative immunoblot analysis of acute brain slices for p35/25 after NMDA/Glycine (100/50 µM, NMDA/Gly) treatment, (n = 3–4 per group) p = 0.0081 Student's t-test. (B) Quantitative immunoblot of acute brain slices for phospho-Thr75 DARPP32 after 1 h treatment 25–106 (10 µM), (n = 4–5 per group) p < 0.0001 Student's t-test. (C) TTC viability staining of acute brain slices subjected to NMDA/Gly (100/50 µM) treatment following preincubation with 25–106, (n = 6–8 per group) F(2,17) = 45.19 p < 0.0001 ANOVA (NMDA vs. NMDA + 25–106, p = 0.0005, Holm-Sidak post hoc). (D) Quantitative immunoblot of Fodrin breakdown products via calpain, p = 0.0062 (top), p = 0.0014 (bottom) Student's t-test (n = 6 per group). (E) Quantitative immunoblot of p25 generation following rTBI, (n = 4 per group) p < 0.0001 Student's t-test. (F) In vivo inhibition of Cdk5 assessed via quantitative immunoblot for P-Ser549/total Synapsin I after treatment with 50 mg/kg 25–106, (n = 6 per group) p = 0.0305 Student's t-test. All data are means ± SEM, *p < 0.05, **P < 0.01, ***P < 0.001 ****P < 0.0001. Full uncropped blots for each panel are provided in Supplemental Fig. 5.

To investigate the role of Cdk5/p25 signaling in rTBI, we assessed the excitotoxic activation of calpain following rTBI. As one index, we examined the breakdown of the calpain reporter, Fodrin into cleaved 150 and 120 kDa products 48 h post-injury55. Rats subjected to rTBI displayed an increased activation of calpain protease (Fig. 5D). This effect corresponded to an increase in the aberrant activation of Cdk5 observed through the production of the excitotoxic cofactor p25 (Fig. 5E). To assess the functionality of 25–106 in vivo, rats were treated with 50 mg/kg I.P. and brain lysates were blotted for Cdk5 dependent phosphorylation states. 25–106 caused a marked reduction in Cdk5 phosphorylation of Synapsin I 6 h after treatment in vivo (Fig. 5F). Together, these studies demonstrate aberrant activation of Cdk5 in excitotoxcity and rTBI and suggest 25–106 may be used as a therapeutic inhibitor of Cdk5.

While conditional knockout of Cdk5 has shown its role in mediating various forms of neuronal injury and neurodegeneration, few Cdk5/p25 specific substrates and mechanisms have been identified. Utilizing this new inhibitor of Cdk5, we investigated the downstream effectors of Cdk5/p25 that may mediate neuronal demise and subsequent cognitive impairments associated with injury through Cdk5 phosphorylation site-directed phosphoproteomics of the hippocampus from control, rTBI, and rTBI rats treated with 25–106 (rTBI + 25–106) 48 h post-injury, which corresponded to the generation of the p25 fragment observed previously (Fig. 6A). For this proteomic analysis, mass spectrometry was conducted on lysates that have undergone an affinity enrichment with antibody-conjugated beads which binds peptides harboring proline-directed consensus phosphorylation sites including those commonly used by Cdk556. Robust changes in the Cdk5 phospho-landscape following rTBI and rTBI + treatment with 25–106 were apparent with 3,021 uniquely modified sites detected via LC/MS–MS (Fig. 6B–E). Rotational head trauma resulted in 653 upregulated phospho-sites with a fold change (FC) ≥ 1.3 (Fig. 6B), and a corresponding decrease in 650 ≤ − 1.3 FC phospho-sites. Interestingly, differentially modified sites included alterations in the phosphorylation of proteins associated with AD such as Microtubule Associated Protein 2 (MAP2), and the Spinocerebellar Ataxia (SCA) associated protein, Diacylglycerol Lipase Alpha (DAGLA)57,58. Additionally, the potential Cdk5-dependent phosphorylation states of proteins associated with cellular demise such as, apoptosis-stimulating protein of p53 (PPP1R13B), and homeostatic synaptic excitability such as GRIN2a (GluN2A) were altered by rTBI (Fig. 6B)59,60.

Cdk5 enriched phosphoproteomics of rTBI. (A) Experimental design schematic displaying tissue isolation, extraction, Cdk5 motif enrichment, and peptide detection and analysis. (B) Volcano plot of differentially phosphorylated proteins between rTBI and con rats. (n = 5 pooled brains in duplicate per group) (increased sites in green decreased sites in red). (C) Volcano plot of differentially phosphorylated proteins between rTBI + 25–106 and rTBI alone rats. (D) Volcano plot of differentially phosphorylated proteins between rTBI + 25–106 and con rats. (E) Heatmap of all differentially phosphorylated sites across three comparisons in B-D expressed by fold change of each phosphor-site. (F) Venn diagram representing overlapping sites with positive FC ≥ 1.3 in rTBI:CON and negative FC in rTBI + 25–106:rTBI. (G) Dot-plot display of Ingenuity Pathway Analysis of phosphoproteins and canonical pathways upregulated FC ≥ 1.3 by rTBI:Con (Top) and downregulated with a FC ≤ − 1.3 by rTBI:Con (bottom). (H) Dot-plot display of Ingenuity Pathway Analysis of phosphoproteins and canonical pathways upregulated FC ≥ 1.3 by rTBI + 25,106:rTBI (Top) and downregulated with a FC ≤ − 1.3 by rTBI + 25–106:rTBI (bottom).

We also observed a reciprocal relationship between the Cdk5 consensus site phospho-landscape of rTBI rats in comparison to rTBI rats treated with 25–106 (Fig. 6E, F), as many phospho-sites that were increased by rTBI were reduced in rTBI rats treated with 25–106. Specifically, of the 653 phospho-sites upregulated by rTBI, 473 of these sites were decreased in the hippocampus of rTBI rats treated with 25–106 representing an inverse relationship of 72.4% of the phospho-sites (Fig. 6F).

Ingenuity Pathway Analysis was used to identify key molecules and interacting pathways evoked by injury (Fig. 6G, H). Upregulated pathways from rTBI highlight molecules linked to neuronal impairments. These included altered phosphorylation states of molecules involved in Ca2+ signaling (Ca2+ Sig.), AMPK signaling (AMPK Sig.), GABA receptor signaling (GABAR Sig.), and protein ubiquitination (Pro. Ub.), as well as pathways invoked in neurodegeneration including amyotrophic lateral sclerosis signaling (ALS Sig) and Huntington's disease signaling (HD Sig.) (Fig. 6G, top). Pathways assessed for proteins in which Cdk5-dependent phosphorylation states decreased following rTBI include glutamate receptor signaling (GluR Sig.), CREB signaling, reelin signaling, and synaptogenesis signaling pathways (SSP), all part of networks necessary for homeostatic neuronal function (Fig. 6G, bottom). Furthermore, the reduced phospho-states following rTBI evoked alterations in molecules associated with neuroinflammatory signaling (Neuroinflam Sig.) and neurovascular coupling signaling pathways (NVCSP) (Fig. 6G). Interestingly, rTBI rats treated with 25–106 (rTBI + 25,106) invoked phosphorylation state increases in pathways such as neurovascular coupling signaling pathway (NVCSP) and synaptogenesis signaling pathways (SSP), both of which displayed a reciprocal relationship in untreated rTBI rats (Fig. 6H). Rats subjected to rTBI + 25–106 also showed many phosphorylation state-dependent increases in proteins associated with inositol phosphate signaling including D-myo-inositol-5-phosphate metabolism (D-m-IP5 met.), 3-phosphoinsitide biosynthesis (3-PI Bs) and degradation (3-PI Deg.), as well as the super pathway of inositol phosphate compounds (SIPC) (Fig. 6H).

Additionally, we observed an inverse relationship in the pathways implicated in the phospho-sites that decreased following rTBI + 25–106. Here, we observed decreased phosphorylation of proteins associated with Huntington's disease signaling (HD Sig.), autophagy, and AMPK signaling, all of which were upregulated by rTBI alone (Fig. 6H). Also, rTBI + 25–106 decreased the phosphorylation states of molecules involved in 14–3-3 signaling including PPAR signaling, IL-15 production, reelin signaling, and SSP compared to TBI alone (Fig. 6H). Compared to controls, rTBI + 25–106 also evoked upregulation of PI3K/AKT, reelin, HIPPO, SSP, AMPK, 3-PI Deg, 3-PI biosynthesis (Bs), and SIPC signaling (Supplemental Fig. 3A). Additionally, compared to controls, rTBI + 25–106 caused downregulation of molecules involved in HIPPO, 14–3-3, GLUR, HD, G-protein coupled receptor, AMPK, Reelin, and SSP signaling (Supplemental Fig. 3B). Altogether, these data demonstrate the Cdk5/p25 signaling in the regulation of various neuronal pathways of injury involved in neuronal plasticity and synapse formation. Interestingly, this Cdk5 site enriched phosphoproteomic library derived from the acute phase of injury also unveiled the regulation of molecules implicated in chronic neurodegenerative conditions such as ALS and HD, two neurodegenerative conditions mediated via Cdk561,62.

As an indication of how the molecules with increased or decreased phosphorylation states alter the biological processes within cells, gene ontology analysis using the open-source platform, Enrichr was employed. The 10 most significant biological processes altered by increased phosphorylation states following rTBI include those highly specified in structural and axonal functions, such as the regulation of microtubule polymerization/depolymerization, axonogenesis, cell morphogenesis in neuronal differentiation, neuronal projection morphogenesis, regulation of supramolecular fiber organization, and cellular component assembly (Supplemental Fig. 3C). This pattern of effects on structural and axonal processes was also observed from gene ontology analysis of downregulated phospho-sites following rTBI including regulation of microtubule polymerization/depolymerization, regulation of supramolecular fiber organization, axon development, cellular component assembly, and cell morphogenesis in neuron differentiation (Supplemental Fig. 3C).

Upregulated phospho-proteins in rats subjected to rTBI and rTBI + 25–106 also displayed similar patterns in the alterations of biological processes within structural and axonal gene ontologies including neuron projection morphogenesis, neuron projection development, cell morphogenesis/neuron differentiation, as well as regulation of microtubule polymerization/depolymerization, regulation of dendritic spine morphogenesis, and synapse organization (Supplemental Fig. 3D). Rats subjected to rTBI + 25–106 also showed decreased phosphorylation in proteins associated with regulation of microtubule polymerization/depolymerization, regulation of supramolecular fiber organization, axon development, axonogenesis, and cell morphogenesis in neuron differentiation relative to rTBI alone animals (Supplemental Fig. 3D). The same pattern of structural and axonal processes was further observed in ontological analysis of rTBI + 25–106 rat hippocampi as compared to controls (Supplement Fig. 3E).

Conditional knockout of Cdk5 activity has been shown to provide neuroprotection from ischemia and experimental models of brain injury. However, the neuroprotective efficacy of pharmacological inhibition of Cdk5 remains unknown23,28. To test the in vivo efficacy of 25–106, rats were subjected to rTBI and treated with 25–106 within 10 min after undergoing the injury procedure. Cognitive recovery was then assessed by fear conditioning 7 days post-injury (Fig. 7A, B). As previously observed, rats subjected to rTBI showed significant reductions in contextual fear learning and memory (freezing behavior). Also, treatment with 25–106 in the absence of rTBI had no effect on basal memory performance. In contrast, rTBI rats treated with 25–106 displayed no significant memory impairment in comparison to uninjured control animals and were significantly improved in mnemonic function compared to animals that received rTBI without Cdk5 inhibition (Fig. 7A). Similarly, rats subjected to rTBI alone showed significant reductions in cued fear learning and memory, while rTBI rats treated with 25–106 displayed cued-leaning and memory rates similar to control animals and significantly improved from rTBI alone (Fig. 7B). Thus, acute post-injury Cdk5 inhibition almost completely blocked cognitive impairment in response to rTBI.

Cdk5 inhibition is neuroprotective against rTBI. (A) Fear conditioning freezing rates in contextual conditioning after rTBI treatment with 25–106 F(3,37) = 4.057, p = 0.0137, ANOVA; (Con-TBI p = 0.0132, TBI-TBI + 25,106, p = 0.0384 Holm-Sidak post hoc) (n = 8–14 per group). (B) Fear conditioning freezing rates in cued conditioning after rTBI treatment with 25–106 F(3,37) = 10.07, p < 0.0001 ANOVA; (Con-TBI p = 0.0002, TBI-TBI + 25,106 p = 0.0137 Holm-Sidak post hoc) (n = 8–14 per group). (C) Immunohistochemistry and quantitation of GFAP+ astrocytes in CA1 following rTBI and treatment with 25–106 F(3,18) = 6.295, p = 0.0041 ANOVA; (Con-TBI, p = 0.0054, Con-TBI + 25,106, p = 0.2259, Holm-Sidak post hoc). (n = 4–6 per group) Scale bars = 50 µm. (D) Immunohistochemistry and quantitation of IBA1+ microglia in CA1 following rTBI and treatment with 25–106 F(3,18) = 8.070, p = 0.0013 ANOVA; (Con-TBI, p = 0.0009, TBI-TBI + 25,106 p = 0.0209, Holm-Sidak post hoc) (n = 4–6 per group) Scale bars = 50 µm. (E) Nissl stained CA1 hippocampal subregion F(3,15) = 14.70 p < 0.0001 ANOVA (CON-TBI, p = 0.0003, TBI-TBI + 25–106 p = 0.01 Holm-Sidak post hoc) Scale bars 20 µm. All data are means ± SEM, *p < 0.05, ** < 0.01, *** < 0.01.

To assess the ability of 25–106 to ameliorate the neuropathological outcomes associated with impaired hippocampal memory, neuroinflammation was assessed via staining for astrogliosis and microgliosis 14 days post-injury (Fig. 7C, D). While rTBI animals alone showed the significant increases in CA1 astrocyte reactivity (GFAP+), 25–106 treatment attenuated this effect. Similarly, 25–106 prevented the neuroinflammatory activation of microglia in the hippocampus CA1 layers (Fig. 7D). In addition to these neuroprotective effects in the CA1 region, increases in gliosis occurred in other hippocampal layers including the dentate gyrus, and CA3 following rTBI and was attenuated by 25–106 (Supplemental Fig. 4A–F). In addition to inflammation, cell death is a major pathological change following TBI. To assess alterations in cell death, neurons displaying chromatolysis were counted. Consistent with other pathological outcomes, rTBI rats displayed an increase in damaged neurons following rTBI. Consistent with the other neuroprotective effects observed, this was ameliorated by treatment with 25–106 (Fig. 7E). Altogether, these data demonstrate the therapeutic potential of systemic Cdk5 inhibition via 25–106 to neuroprotect from rTBI.

Rotational TBI remains the most clinically relevant form of brain trauma63. However, understanding of these injuries has been, limited by the lack of rotational acceleration-induced injury models in rodents that accurately recapitulate patient phenotypes. Specifically, most preclinical rotational head trauma has been limited to large animal models due the conserved gyrencephalic brain structures lacking in rodent models64. While these large animal models of brain injury offer some physiologically relevant advantages, they remain costly and underpowered for genetic manipulations and preclinical screening of therapeutics65. The fundamental challenge with rodent models of rTBI has been to scale rotational forces to small brain structures. Helmet telemetry data in humans have determined a typical sports-related head trauma induces rotational accelerations between 1 and 3 Krad/s230,66. Scaling to the 2 g rat brain, these equate to rotational forces greater than 350 Krad/s231,67.

Previously, rodent models including the Medical College of Wisconsin (MCW) and Closed-Head Impact Model of Engineered Rotational Acceleration (CHIMERA) have provided insights into modeling these injuries in small mammals. These models have demonstrated histopathological and behavioral abnormalities following injury and show differential outcomes in these injury models in comparison to blast models of TBI68,69,70. However, these models have remained limited largely to initial characterizations and, in some instances, lack equivalent scalar angular forces. Here, we validate our rTBI model's rotational kinetics and demonstrate the feasibility for recapitulating numerous pathophysiological outcomes of head injuries across post-injury delays. Importantly, we have demonstrated several aspects of injury with this novel model of repetitive rTBI (summarized in Table 1). However, each of these outcomes depend upon various starting input conditions such as impact severity, animal age, anesthesia conditions, and post injury delay. Determining how each input variable effects the exact thresholds and temporal regulation of each pathological change observed here remains an important question.

Several animal studies indicate neuroinflammation as a key TBI consequence18. Depletion of infiltrating cells and halted neuroimmune responses improve behavioral and histological outcomes following cortical impact or fluid percussion injury in rodents78,79. Similarly, we show our rTBI model induces diffuse neuroinflammation. This inflammation is observed acutely (48 h) and persists into longer post-injury delay periods (7 and 14 d). We employed in vivo PET/CT imaging of TSPO as a potentially translational non-invasive diagnostic for rTBI. Increased TSPO expression likely indicates production of reactive oxygen species within neuroimmune cell populations such as microglia and astrocytes. This marker has been used clinically in patients with neurodegenerative conditions and in former professional football players exposed to chronic head injuries. The use of this marker as a diagnostic for acute brain injury in patients remains understudied32,34. Interestingly, we observed no alteration in global brain uptake of TSPO, suggesting a mild or moderate neuroinflammatory phenotype. However, significant alterations in TSPO uptake were present when examining specific brain regions. The significance of region-specific neuroinflammatory responses such as these and the overall diagnostic sensitivity of this noninvasive technique patients with rTBI remains an important translational question.

Additionally, we observed the presence of increased hyperphosphorylated Tau in the form of AT8 immunoreactivity throughout the brains of rats subjected to rTBI 6 months post-injury. AT8 staining is specific for Tau that is abnormally phosphorylated at both Ser202 and Thr205, and the presence of AT8 Tau inclusions represents an underlying pathology in the post-mortem diagnosis of CTE38. However, the preclinical understanding and development of CTE therapies remains limited by our abilities to model this disease preclinically. While some TBI models have shown acute increases in phosphorylated Tau, limited studies have demonstrated Tau inclusions and other CTE-like pathologies80. In most studies, researchers have relied on overexpression of human isoforms of Tau and demonstrating an increased potential to aggregate following head trauma80. While our finding here is exciting, more work will need to be conducted to determine the chronic effects of rTBI and the corresponding CTE-like pathologies including: the degree of TDP-43 pathology, stereological assessments of depleted neuronal populations, and solubility/isotypes of Tau inclusions. Notably, we did not observe increases in Tau phosphorylation at earlier stages of injury (14 days post-injury) via immunoblotting (data not shown). Assessing the temporal regulation of Tau phosphorylation remains an interesting question within this model and represents exciting potential for future research.

Brain injury results in a multitude of cognitive and behavioral impairments. Some of the most common symptoms reported by TBI survivors are persistent impairments in cognitive abilities including deficits in memory, attention, and executive functioning8,42. These features are commonly linked to structural damage in the hippocampal formation following injury37,81, and numerous animal models point to impaired hippocampal-mediated contextual and spatial memory impairments following TBI35,50,70. In agreement, our rTBI model caused impaired hippocampal-dependent contextual fear memory and correlated with neuroinflammation in this region. Interestingly, amygdala-dependent cued fear memory was also impaired and exhibited a neuroinflammatory response. Structural damage to these areas, as well as brain-wide diffuse axonal injuries including interconnected efferent and afferent networks of the hippocampus/amygdala may be causally linked to these effects.

Underlying the observed memory impairment, rTBI caused an almost complete ablation of hippocampal long-term synaptic plasticity. This is consistent with the electrophysiological effects of other TBI models including fluid percussion, controlled cortical impact, and blast which also cause circuitry damage within the interconnected layers of the hippocampus82,83,84. These alterations in hippocampal LTP occur in acute phases and persist into more chronic stages of TBI23,36,50,84. However, the long-lasting effects of rTBI on LTP remain to be further characterized. While these alterations in axonal function likely link to axonal damage and structural changes in synaptic connections, assessing brain connectivity and structural alteration within axons remains an exciting question. TBI pathophysiology involves cellular depolarization, voltage-gated Ca2+ channel activation, massive glutamate release, and activation of Ca2+ -dependent proteases such as calpain85,86. Calpain represents a potential therapeutic target for TBI87 and preclinical studies show promising calpain inhibitor neuroprotective activity88,89,90,91. Cdk5/p25 activation is a principle downstream effector of excitotoxic calpain activation. Invocation of aberrant Cdk5 is broadly implicated in neuronal injury and neurodegeneration88,89,90,91,92,93,94,95,96. Conditional knockout of Cdk5 is neuroprotective in models of ischemia and experimental brain injury23,28. Transgenic overexpression of the aberrant co-activator of Cdk5, p25, induces rapid onset of AD in mice models52. Inhibition of p25 activity using small interfering peptides is neuroprotective in PD models97. Therefore, aberrant Cdk5 activity represents a critical therapeutic target for acute injury and neurodegenerative conditions.

Given the potential efficacy of Cdk5 inhibition to block the deleterious behavioral and neuroinflammatory effects of rTBI, we used phosphoproteomics to enrich for phosphopeptides derived from potential Cdk5 substrates in the hippocampus. We report hundreds of novel potential Cdk5/p25 substrates phosphorylated in response rTBI but blocked when injury was followed by 25–106 treatment. Bioinformatic analysis implicated many of these in various cascades associated with cellular demise including AMPK, Ca2+, and neurodegenerative signaling. Importantly, human TBI survivors have a 56% increased risk of PD and a 2.3 times increased risk of developing AD21,98. Since Cdk5/p25 activity is associated with the progression of both of these neurodegenerative conditions, it is likely that some of these substrates identified in the acute phases of rTBI likely contribute to longer lasting neuropathological effects of neurodegeneration. Understanding the dynamics of these phospho-substrates in the progression of TBI and as potential biomarkers of future brain degenerative conditions remains an exciting and expansive question generated by this work.

Several specific results within the global proteomic dataset derived here are also consistent with both preclinical models and clinical studies. For example, total proteome analysis in both rTBI and rTBI + 25–106 showed the highest upregulation of bradykinin family members Kng1 (FC = 4.0, rTBI FC = 6.6 rTBI + 25–106) and Kng2 (FC = 3.8, rTBI FC = 6.0 rTBI + 25–106). These proteins have been heavily implicated in animal TBI models as well as clinical studies99. Kng1/Kng2 serve as ligands for B2 bradykinin receptors. In alignment with these targets as mediators of rTBI, B2 receptor antagonists have been tested in clinical trials for TBI treatment100. We note that the induction of this inflammatory signaling cascade was not responsive to 25–106, and thus may exemplify mechanisms of injury that are not necessarily effectors of aberrant Cdk5 activation.

There is currently no approved treatment to ameliorate the neuropathological consequences of TBI. Having recently discovered the first brain-active systemic Cdk5 inhibitor, 25–10654, we tested it here as a possible treatment for rTBI. While use of 25–106 in the acute phase of rTBI appears promising, injury delay periods to emergency room treatments can vary between patients and nearly 80% of TBI patients are initially treated and then released from emergency departments1. The efficiency of Cdk5 inhibition in more sub-acute and chronic stages of rTBI remains to be explored. Current TBI patient treatments are directed at symptom management including anticonvulsants for seizures, diuretics to reduce pressure in the brain, and stimulants and anti-anxiety medications to increase alertness and reduce negative mood states101,102,103,104. Clinical trials have investigated numerous possible neuroprotective interventions including NMDA antagonists to reduce excitotoxicity and anti-inflammatory agents but have not significantly improved outcomes100,105. On the other hand, clotting agents such as tranexamic acid can prevent brain edema in the acute phase, although this treatment is limited to intracranial hemorrhages and may not be a useful approach for repetitive mild injuries mediated by axonal injury and inflammation106. Altogether, this work brings forth a new model to the field of brain injury, assesses a novel neuroprotective treatment, and progresses our knowledge of the mechanisms mediating neuronal death caused by head trauma and other excitotoxic conditions.

All methods were performed in accordance with relevant guidelines and regulations for safety and animal welfare of the University of Alabama at Birmingham (UAB) and approved by UAB. This study is in accordance with the ARRIVE guidelines.

Male Sprague Dawley rats (9–10 months old, Envigo) were used for all TBI experiments. All were single housed and maintained on a normal 12 h day/night cycle. For euthanasia in biochemical experiments, brains were collected via rapid live decapitation in the absence of anesthesia. For histological experiments, rats were euthanized via intraperitoneal injections of an anesthetic mixture of (100 mg/kg) ketamine (10 mg/kg) xylazine. For rTBI procedures, rats were maintained under anesthesia (isoflurane) at 1.5% for the duration of the procedure. All experiments were performed under approved protocols by the University of Alabama at Birmingham (UAB) Institutional Animal Care and Use Committees (IACUC).

All rats were habituated to the room in which brain injuries occurred for 1 h, anesthetized using 1.5% isoflurane vapor, and sedation was maintained throughout the procedure via a nose-cone within the subject's helmet. Anesthetized rats were placed in a custom designed harness that was inserted into the base of the pendulum and locked tightly into place. The rat head was fitted into a freely rotational helmet with foam padding to secure head placement and rotation in one direction. Flowing N2 gas was allowed to fill an Arduino controlled closed solenoid value connected to an air motor and the pendulum was adjusted to a 90˚ angle raised approximately 84 cm from the base of the machine. Launch was electronically executed using custom software (LabVIEW) to open the solenoid allowing N2 to propel the pendulum forward towards the ventral strike plate. Helmet impact results in a 90˚ rotation of the helmet and animal's head in both directions. Rotational forces were derived in LabVIEW from helmet velocity and exported into Excel. Each animal received 10 repetitive impact rotations over a 30 min period. Controls were maintained in the machine, under anesthesia for 30 min with no head rotation. Following injury each animal was assessed for normal locomotion and righting reflex abilities.

Noninvasive positron emission tomography-computed tomography (PET/CT) imaging was conducted 7 days post TBI with [18F]DPA-714, a translocator protein (TSPO) radioligand produced at the Cyclotron Facility at the University of Alabama at Birmingham. Rats were injected with 300 µCi (11.1 MBq) of [18F]DPA-714 intravenously. Rats were immediately scanned using a GNEXT small animal PET/CT (Sofie Biosciences) for 30 min. Imaged regions of interests (ROIs) within the brain were drawn with CT guidance using VivoQuant (Invicro) and a 3D rat brain atlas (Allen brain atlas) was applied to automatically segment each brain region. The mean, maximum, hotspot, and heterogeneity of standard uptake values (SUVs) were determined using the formula: SUV = [(MBq/mL) × (animal wt. (g))/injected dose (MBq)].

Histology was performed as described107. Briefly, rats were transcardially perfused with 1X PBS/50 mM NaF followed by 10% formalin fixation. Brains were submerged in 10% formalin and fixed overnight, paraffin embedded and serially sectioned at 5 µm at Bregma level -3.3 to -4.5. Sections were deparaffinized in 100% xylene 3 × 5 min. Following deparaffination, slides were rehydrated using ethanol gradients of 5 min incubations in 100%EtOH, followed by 95%EtOH, and lastly 75%EtOH before being dipped in H2O for 1 min. Following rehydration, slides were incubated in 1 × pre-warmed citrate antigen retrieval buffer (Thermo Fisher) and heated for 10 min in a pressure cooker followed by 20 min of cooling in the citrate buffer. Sections were then permeabilized and blocked in 0.03% Trition X-100/PBS containing 3% goat serum. DAB-stained sections underwent a 10 min incubation in 10% horseradish peroxidase. Immunohistochemistry was performed using glial fibrillary acidic protein (GFAP) (1:1000; Millipore) and ionized Ca2+-binding adapter protein (1:1000; Wako) incubated in 0.3% Tween 20/PBS overnight. Following 1 h of PBS washes, fluorescent visualization was performed using secondary antibodies Cy3 anti-rat (1:500), Alexa 488 anti-mouse (1:200) (Jackson Immuno) and imaged using an Olympus BX60 microscope mounted with Olympus DP74 digital camera. DAB-stained sections were incubated with anti-mouse biotin-conjugated secondary (1:500) or anti-rabbit biotin-conjugated secondary (1:500) (Jackson Immuno) followed by PBS washing and 30 min incubation with ready-to-use streptavidin peroxidase. DAB staining was performed using DAB substrate Kit (abcam) according to manufacturer's instructions and sections were incubated until brown precipitate was visible (5 min, AT8; 1 min SMI-31). Following DAB staining sections were incubated in hematoxylin counterstain and for 1 min, washed with H2O, dehydrated in 100% EtOH (3 × 5 min) and cleared in xylene (3 × 5 min). For Nissl stains, paraffin embedded sections were deparaffinized and rehydrated in serial EtOH washes (as described above). Slides were then incubated in warmed (50 °C) 0.1% cresyl violet solution (Electron Microscopy Sciences) for 10 min. Following staining Nissl sections were differentiated in 95% EtOH, dehydrated in 100% EtOH, and cleared with 100% Xylene. All stains were performed using a minimum of 3 sections per animal and 4–6 animals per group. All stains within groups were analyzed using the same background thresholds (0–72; GFAP), (4–110; IBA1), and (72–234; AT8). Following thresholding, percent area of immunofluorescent positive staining was determined using ImageJ particle detection software (Fiji), optical density analysis of DAB stained sections was performed as described108 and OD was determined by log (255/mean intensity) for 8bit images. Dark Nissl-stained neurons were detected using (Fiji) multipoint counting tool.

Brain slice pharmacology was performed as described54. Briefly, brains were rapidly decapitated and submerged in ice cold Normal Kreb's solution (125 mM NaCl, 2.5 mM KCl, 1.25 mM NaH2PO4, 25 mM NaHCO3, 1.1 mM MgCl2, 2 mM CaCl2 and 25 mM glucose). Brains were coronally sectioned at 350 µm using a vibratome in regions of interest. Slices were recovered in oxygenated Krebs solution at 30˚C. Following recovery, slices where incubated in Kreb's containing pharmacological interventions indicated of NMDA/Glycine (100 µM NMDA, 50 µM Gly, 1 h), 25–106 (10 µM, 1 h). Following treatment, slices were snap frozen in dry ice to terminate treatment. Following NMDA treatment, slices were incubated in 0.125% 2,3,5-Triphenyltetrazolium chloride (TTC) viability stain for 20 min. Stained slices were fixed in 4% PFA for 10 min and scanned. TTC viability was assessed by mean intensity of staining and expressed as percentage of control viability.

Immunoblotting was performed as previously described107. Briefly, rat brains were rapidly dissected and submerged in a 4˚C solution of PBS/50 mM NaF. Subregions were dissected and snap frozen. Brain regions were subsequently homogenized in 1% SDS/50 mM NaF and sonicated at 40 dB pulses until tissue was completely homogenized. Protein concentrations were determined from lysates via BCA protein assay. Samples were diluted in 4 × lysis buffer and proteins were separated by molecular weight via SDS-PAGE. Proteins were then transferred to 0.45 nm nitrocellulose, blocked in Licor Blocking Buffer, and incubated with 1° antibody (Ab) overnight. Membranes were washed with 1xTBS-T and incubated with Licor fluorescent 2° Ab for 1 h at room temperature. Proteins expression was visualized using Licor Odyssey CLx membrane scanner. Arbitrary units of florescent intensity for each protein band was quantified using ImageStudio. Phospho-band intensity was normalized to total protein bands, total protein bands were normalized to actin loading controls, and cleavage products were normalized to un-cleaved total protein. Antibodies used include p35/p25 (Cell Signaling Technology), phospho-Ser549 and total Synapsin I (PhosphoSolutions), Fodrin (Enzo Life Sciences), phospho-Thr75- and total DARPP32 (Cell Signaling Technology), EPHx2 (Abcam), Legumain (Cell Signaling), AT8 (Invitrogen), Actin (Invitrogen).

Neurophysiological studies were conducted in rats 7 days post-injury as described50. Briefly, brains were rapidly dissected in ice-cold artificial cerebrospinal fluid (ACSF; 75 mM sucrose, 87 mM NaCl, 2.5 mM KCl, 1.25 mM NaH2PO4, 25 mM NaHCO3, 7 mM MgCl2, 0.5 mM CaCl2 and 10 mM glucose). Transverse hippocampal slices (350 μ) were sectioned using a vibratome (Leica Microsystems Inc., VT1000S) in NMDG cutting/recovery solution (N-methyl D-glucamine (100 mM), KCl (2.5 mM), NaH2PO4 (1.2 mM), NaHCO3 (30 mM), HEPES (20 mM), MgS04 (1 0 mM), CaCl2 (0.5 mM), and glucose (25 mM) at 30 °C (pH 7.3–7.4). After 2 min, slices were transferred to HEPES holding solution NaCl (92 mM), KCl (2.5 mM), NaH2CO3 (30 mM), NaH2PO4 (1 mM), HEPES (20 mM), D-Glucose (25 mM), MgCl2 (1 mM), CaCl2 (1 mM) for 1 h at 30 °C. Slices were allowed to incubate for 30 min in recording solution of oxygenated Kreb's (125 mM NaCl, 2.5 mM KCl, 1.25 mM NaH2PO4, 25 mM NaHCO3, 1.1 mM MgCl2, 2 mM CaCl2 and 25 mM glucose) prior to recording.

Recordings were performed using a Multiclamp 700A amplifier with a Digidata 1322 and pClamp 10 software (Axon, Molecular devices, LLC). Field excitatory postsynaptic potentials (fEPSP) from CA1 were evoked by square current pulses (0.1 ms) at 0.033 Hz with a bipolar stimulation electrode (FHC, Bowdoinham, ME). Stimulus intensity was defined using a stimulus intensity required to induce 50% of the maximum EPSP slope using the input–output curves. The sample intensity was used for PPR recordings across different intervals. A stable baseline was recorded for at least 10 min prior to high frequency stimulation (HFS, 4 trains, 100 Hz, 1 s duration, separated by 20 s). Post-tetanic potentiation (PTP) was analyzed by taking the average of the slopes from the traces recorded during the first 2 min after HFS. LTP was assessed for at least 45–50 min following HFS. The PPR values were calculated by dividing the second fEPSP slope by the first fEPSP slope (fEPSP2/fEPSP1). All recordings were performed in the absence of any drug treatment and only 1 or 2 slices were recorded from each individual rat. Data were analyzed with Clampfit 10 software (Axon, Molecular devices, LLC).

Fear conditioning studies were performed as described109. Briefly, rats were habituated to the behavioral room for 1 h before experimentation. Rats were placed in fear conditioning chambers (Med Associates) to establish baseline freezing rates between cohorts. The following day rats were subjected to fear conditioning training. Each rat was allowed to freely explore the chamber for 2 min followed by a 30 s tone terminating in a mild foot shock (0.7 mA). Rats remained in the chamber 2 min after shocking before returning to their home cage.

Context-dependent fear memory was assessed 24 h post-shock training. Rats were re-introduced into the conditioning box for 5 min. Freezing responses (motionless except respirations) were recorded using VideoFreeze software. Cued fear conditioning memory was assessed 27 h post-shock training where rats were allowed to explore a novel context with novel odor (vanilla). The rats were left exploring the novel context for 3 min without tone followed by a 3 min period with the training tone playing. Freezing responses (motionless except respirations) were recorded.

Shock sensitivity and nociceptive responses were assessed by returning rats to fear conditioning chambers and evaluating minimal thresholds to induce rat flinching, jumping, and vocalization of pain across adverse stimuli (0–1.5 mA) shocks.

Freshly dissected and snap frozen rat hippocampi were sonicated, centrifuged, reduced with DTT, and alkylated with iodoacetamide. Total protein for each sample (10 mg) was trypsin digested, purified over C18 columns (Waters), enriched using the PTMScan Phospho CDK + CDK/MAPK Substrate Motif Antibodies (#9477/#2325 Cell Signaling Technology) and purified over C18 tips as previously described110. For total proteome analysis an additional 100 mg of each sample was digested with LysC and trypsin and digested samples were purified over C18 tips, labeled with TMT 10-plex reagent (Thermo), bRP fractionated (96 fractions concatenated non-sequentially to 12), and C18 purified for LC–MS/MS as previously described111. LC–MS/MS analysis was performed using an Orbitrap-Fusion Lumos Tribrid mass spectrometer as described110,111 with replicate injections of each sample run non-sequentially for the phosphopeptide analysis. Briefly, peptides were separated using a 50 cm × 100 µM PicoFrit capillary column packed with C18 reversed-phase resin and eluted with a 90 min (Phospho) or 150 min (TMT total proteome) linear gradient of acetonitrile in 0.125% formic acid delivered at 280 nl/min. Full MS parameter settings are available upon request. MS spectra were evaluated by Cell Signaling Technology using Comet and the GFY-Core platform (Harvard University)112,113,114. Searches were performed against the most recent update of the NCBI Rattus norvegicus database with a mass accuracy of ± 20 ppm for precursor ions and 0.02 Da product ions.

Results were filtered to a 1% peptide-level FDR with mass accuracy ± 5 ppm on precursor ions and presence of a phosphorylated residue for CDK Substrate enriched samples. TMT total proteome results were further filtered to a 1% protein level false discovery rate. Site localization confidence was determined using AScore115. All CDK Substrate quantitative results were generated using Skyline116 to extract the integrated peak area of the corresponding peptide assignments or in GFY-Core using signal: noise values for each peptide and summing individual signal: noise values for all peptides for a given protein for TMT total proteome data. Accuracy of quantitative data was ensured by manual review in Skyline or in the ion chromatogram files. Quantitative data was normalized across samples using median abundance for CDK Substrate data or sum signal: noise for TMT total proteome data.

Differentially modified phosphoproteins across all groups upregulated ≥ 1.3-fold change or downregulated ≤ − 1.3 -fold change were subjected to Ingenuity Pathway Analysis (IPA), (Qiagen). Log-transformed p-values of significantly enriched canonical pathways and the phosphorylated/dephosphorylated molecules comprising each pathway were used to construct dot plots of canonical pathways significantly Up/downregulated in each condition. IPA Z-scores were used to predict activation or inhibition of each canonical pathway. Phosphoproteins exhibiting fold changes of greater ≥ 1.3 or ≤ − 1.3 were subjected to gene ontology enrichment analysis via open source gene enrichment analysis software, Enrichr117. Biological processes and molecular functions from each gene list were derived based and the top 10 processes/functions were derived based on adjusted p-values.

Prior to analysis, all data was examined for normalcy using the Shapiro–Wilk test. In cases of parametric data with normal distributions of two group means Student's t-test were used. In cases of non-parametric comparison of means, the Mann–Whitney test was employed. When comparing more than two group means, one or two-way ANOVAs were used with Holm-Sidak post-hoc tests. When comparing two experimental variables within the same animal, Two-way repeated measures ANOVA was used. For all data, * = p < 0.05, ** = p < 0.01, ** = p < 0.001, *** = p < 0.0001. All statistical analysis was performed using Prism 6 (GraphPad Software, Inc.).

All datasets used in this study are available from the corresponding author upon reasonable request.

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We thank the UAB Behavioral Assessment Core for assistance, and Carlo Wayan, Curtis Anderson, Jose Becerra, Stacy Donovan, and Athanasios Spiropoulos at UT Dallas Mechanical Engineering Senior Design Program. We thank Divya Shaw and the UT Arlington Mechanical and Aerospace Engineering Dept., and Naveen Varma and the UAB School of Engineering. We thank V. Yang for assistance with data library deposition. We thank the UAB Comprehensive Cancer Center's Preclinical Imaging Shared Facility (P30CA013148). We thank University of Texas Southwestern Dept of Psychiatry and Carol Tamminga for support of this work. We appreciate the IACUC staffs at UTSW and UAB for help in developing this model.

This research was facilitated by NINDS T32 Predoctoral Training Fellowship (NS061788) (A.U.). Aspects were supported by pilot funding from the Yale NIDA Neuroproteomics Center (DA018343, A.C.), the DOE (DE-NA 0003962, H.L.) and the University of Texas Louis A. Beachel Jr. endowed Chair (H.L.). This work was also supported by UAB Radiology/CCTS Pilot Research Award. Model development was supported by the Texas Institute for Brain Injury and Repair. Aspects of this work were facilitated by NIH grants NS073855, MH116896, and MH126948 (J.A.B). Aspects of this work were also facilitated by an SDHB Pheo Para Coalition Investigator Award, and an NETRF Accelerator Award.

Department of Surgery, University of Alabama at Birmingham, Birmingham, AL, USA

Alan Umfress, Suma Priya Sudarsana Devi, Raegan Adams, Daniel Epstein & M. Shahid Mukhtar

Department of Translational Neuroscience, University of Arizona College of Medicine in Phoeni, Biomedical Sciences Partnership Bldg, Phoenix, AZ, 85004 , USA

Ayanabha Chakraborti & James A. Bibb

Department of Radiology, University of Alabama at Birmingham, Birmingham, AL, USA

Adriana Massicano & Anna Sorace

Eppley Institute for Research in Cancer and Allied Diseases University of Nebraska Medical Center, Omaha, NE, USA

Sarbjit Singh & Amarnath Natarajan

Department of Pharmacology and Toxicology, University of Alabama at Birmingham, Birmingham, AL, USA

M. Iqbal Hossian & Shaida A. Andrabi

Department of Genetics, University of Alabama at Birmingham, Birmingham, AL, USA

David K. Crossman

Department of Biology, University of Alabama at Birmingham, Birmingham, AL, USA

Nilesh Kumar & M. Shahid Mukhtar

Karagozian & Case, Inc., Glendale, CA, USA

Huiyang Luo

Cell Signaling Technology, Danvers, MA, USA

Claire Simpson, Kathryn Abell, Matthew Stokes & Thorsten Wiederhold

OSF Healthcare Illinois Neurological Institute, Peoria, IL, USA

Charles Rosen

Department of Mechanical Engineering, University of Texas at Dallas, Dallas, TX, USA

Hongbing Lu

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A.U., A.C., S.P.S.D., D.E., R.A., A.M., A.S., I.H., S.A., T.W., C.S., K.A., and M.S. conducted experiments, collected data, and performed formal analysis. H. Luo and H. Lu. engineered the model. S.S. and A.N. synthesized reagents. D.C., N.K., and S.M. conducted analysis and data visualization. A.U., C.R., J.A.B. conceived the study and experimental design. All authors contributed to editing and approval of the final manuscript.

Correspondence to James A. Bibb.

The authors declare no competing interests.

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Umfress, A., Chakraborti, A., Priya Sudarsana Devi, S. et al. Cdk5 mediates rotational force-induced brain injury. Sci Rep 13, 3394 (2023). https://doi.org/10.1038/s41598-023-29322-4

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Received: 27 September 2022

Accepted: 02 February 2023

Published: 28 February 2023

DOI: https://doi.org/10.1038/s41598-023-29322-4

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