Shockwave generation and propagation

A custom-built, compression-driven ‘Schedule 80’ shock tube was used to generate high-pressure, which describe the overpressure that occurs following a real-life blast that can lead to a blast-induced TBI. To make this arrangement successful, the main setup included a 3/4-inch inner diameter, a 4-inch driver, and a 30-inch driven section separated by a Dead Soft aluminum diaphragm [28], which isolated the driver from the driven region containing APHCs that were placed at the end of the driven section. Shockwaves were generated by filling the driver with compressed air until the diaphragm ruptured, sending a shockwave through the driven section (see Fig 1B). Cells were exposed under four conditions: two high blast overpressure levels—single (14.5 psi / 100 kPa for 1.76 ms, classified as mild severity) and double (29 psi / 200 kPa for 3.52 ms, classified as moderate severity) based on established blast injury severity thresholds [29], and two directions— ‘top-to-bottom’ (simulating downward-directed blast forces) and ‘bottom-to-top’ (simulating upward-directed blast forces). In the ‘top-to-bottom’ setup, the shockwave passed through the lid of the petri-dish containing the APHCs, air, cell media, and then the cells. In the bottom-to-top setup, the shockwave was applied directly to the base of the petridish for a more direct impact (Fig 1C). Since wave impedance is a function of material density and wave speed, a transition between materials with different impedance alters the energy distribution, determining the reflected and transmitted wave intensities. Due to these abrupt transitions in pressure, shockwaves can significantly influence cellular behavior and interactions. For example, in the ‘top-to-bottom’ setup, each layer: lid, air, media that introduces an impedance mismatch, reflects part of the wave, weakening the transmitted shockwave and minimizing cytoskeletal disruption. This relationship between reflection and transmission at each interface is described by Zhao et al. [30] as:

(1)

(2)

(3)

where denotes the wave impedance ratio, with and as the densities of materials 1 and 2, respectively, and C₁ and C₂ as the corresponding wave speeds in each material.

Multiple transitions dissipate energy at each interface, the ‘top-to-bottom’ shockwave loses strength before reaching the cells. In contrast, ‘bottom-to-top’ exposure reduces loss of energy, enhancing shear forces, and producing stronger shockwaves. Thus, impedance matching can control how much of the shockwave will be reflected or transmitted at each interface during shockwave propagation across different materials.

Cell culture

Adult rat hippocampal progenitor cells (AHPCs) were gifted by Dr. F.H. Gage, Salk Institute, La Jolla, CA. The AHPCs were isolated from the hippocampus of adult Fisher 344 rats with prior approval from the Animal Care and Use Committee of the Salk Institute. AHPCs were cultured in a T25 tissue culture flask (Thermo Fisher Scientific) coated with poly-L-ornithine (10 µg/mL; Sigma Aldrich) overnight at room temperature, rinsed with an excess volume of phosphate buffered saline (PBS) the following day, and incubated with a laminin (10 µg/mL; Cultrex by Trevigen) solution made in PBS. AHPCs were then incubated overnight at 37° C in 5% CO₂ incubator. The following day the laminin solution was removed, rinsed with PBS and stored frozen until use. Cells were cultured in maintenance medium (MM) composed of Dulbecco’s modified Eagle’s medium/Ham’s F-12 (DMEM/F-12, 1:1; Gibco by Thermo Fisher Scientific) supplemented with 2.5 mM GlutaMAX (Thermo Fisher Scientific), 1 x N2 supplement (Gibco by Thermo Fisher Scientific),1 x Penicillin/streptomycin (Gibco by Thermo Fisher Scientific), and 20 ng/mL basic fibroblast growth factor (bFGF; Gibco by Thermo Fisher Scientific). The cells were incubated at 37 °C in a 5% atmosphere. AHPCs in the flask were fed with MM every 2 days by performing half media changes. They were monitored daily to ensure they were contamination-free and harvested when the cell confluency reached 80%. The multipotential nature of the AHPCs was verified by immunocytochemistry.

Atomic force microscope (AFM) setup

Nanomechanical measurements of the AHPCs were performed using an atomic force microscope (AFM) (Bruker BioScope Resolve system) integrated with a NanoScope V controller. The AFM was operated in Peak Force Quantitative Nanomechanical Mapping (PF-QNM) mode, which simultaneously provided high-resolution imaging and nanomechanical property mapping. This mode provided precise measurements of key mechanical parameters such as Young’s modulus, deformation, dissipation, and adhesion, all while minimizing the risk of damaging the delicate biological samples. Cells were cultured for 72 h under standard growth conditions to reach the desired confluency and were then exposed to single or double blast-induced shockwaves at defined pressure levels using the shock tube apparatus, as detailed in the shockwave exposure protocol. Immediately after shockwave exposure, the dish with AHPCs was placed on the AFM stage (connected to the sample heater) in a vibration isolation chamber to maintain a controlled temperature environment during measurements. Control cells were also retrieved from the incubator alongside treated cells to maintain consistency for comparison. Pre-calibrated PFQNM-LC-CAL-A probes, specifically designed for live cell applications, were employed for single-cell measurements. These probes, with a 70 nm tip radius and a nominal spring constant ranging from 0.074 N/m to 0.101 N/m, offered the necessary sensitivity and flexibility for probing soft biological materials without inducing excessive deformation. A controlled force of 300 pN was applied during each indentation, ensuring gentle yet consistent contact with the cell membrane and minimizing structural disruption. The AFM scan rate and scan size were set at 0.242 Hz and ranging from 17 m to 50 m, respectively, with 256x256 pixel size, achieving a balance between spatial resolution and throughput. This allowed for accurate nanomechanical mapping of each cell’s surface topography, capturing critical mechanical properties. For viscoelastic property measurements, we utilized the AFM ‘Ramp and Hold’ to maintain a constant distance between the AFM tip and the cell while tracking time-dependent force. We held the probe on the cells for 2 seconds and captured these stress relaxation curves using an MLCT-D probe in contact mode. These probes featured a tip radius of 20 nm and a spring constant of 0.03 N/m, making them suitable for capturing the time-dependent mechanical behavior of the cells. All measurements were performed under carefully controlled ambient conditions, with temperature and humidity regulated to preserve cell viability throughout the experimental process.

Analysis for nanomechanical measurements

The nanomechanical data obtained from individual cells were processed and analyzed using Bruker NanoScope Analysis software. This software enabled a detailed examination of both the structural and mechanical properties of the cells at the nanoscale. High-resolution images and nanomechanical maps were exported as .spm files, providing comprehensive visualization of cell surface topography and mechanical characteristics. Force curves were captured as .pfc files at 10 random locations on each cell and analyzed using the Hertz model [31,32] with a spherical indenter, yielding average Young’s modulus values for 20 cells to assess their stiffness and elastic properties. In the Hertz model, the indentation force is written as follows:

(4)

Indentation force F, Young’s modulus E, Poisson ratio , indentation , and R representing the radius of the indenter are the parameters included in the Hertz model.

To characterize viscoelastic behavior, time-dependent force curves were obtained by maintaining a fixed position of the height sensor for two seconds during each measurement. The resulting stress relaxation curves were analyzed using Kohlrausch-Williams-Watts (KWW) models [33] to extract parameters such as retardation time and viscosity described by the following expression:

(5)

The final stress is represented by as time (t) goes towards infinity. Likewise, is the function representing stress which is dependent on t. The shape parameter () and characteristic life () have relationships with the time that the load has been applied to. These metrics characterize the viscoelastic behavior of the material and give insight into how the stress develops when the load has been applied for a very long period of time. A total of 40 data points per cell type were analyzed, offering a thorough assessment of the viscoelastic properties of the cells. Representative stress relaxation curves for each condition are shown in Fig 4, illustrating the distinct mechanical responses of the different cell lines.

Propidium iodide (PI) staining

Cell viability was determined using propidium iodide (PI; Sigma Aldrich) to stain dead cells within the cultures. For the ‘top-to-bottom’ shockwave exposure experiments, PI staining was performed at 5 days in vitro (DIV) (24 hours after shockwave blast injury) and at the end of the culture period (S1 Fig A-B). For the ‘bottom-to-top’ shockwave exposure experiments, PI staining was performed within 6 hours following shockwave exposure and at the end of the culture period (S1 Fig C). The PI was diluted to 1.5 µM in cell culture medium. The cells were incubated in PI solution for 20 minutes at 37°C in 5% CO₂. As a reagent control, one coverslip with cells was incubated with 70% ethanol for 5 minutes to intentionally kill all the cells prior to the addition of the PI solution. After the 20-minute incubation, the cells were rinsed with ice-cold 0.1 M PO₄ buffer for one minute and then were fixed with 4% paraformaldehyde (PFA, Thermo Fisher Scientific) made in 0.1 M PO₄ for 20 minutes. Cells were then rinsed three times for 7 minutes each with phosphate-buffered saline (PBS) to remove all the fixative and then incubated with 4^′,6-diamidino-2-phenylindole (DAPI; 1:500; Invitrogen by Thermo Fisher Scientific) diluted in PBS for 60 minutes at room temperature in the dark. The cells were then rinsed four times for 8 minutes each with PBS. Coverslips were then mounted onto microscope slides with Fluoromount-G with DAPI (Invitrogen by Thermo Fisher Scientific). Propidium iodide staining was used to evaluate cell survival. As a positive control for the PI reagent, samples with AHPCs were incubated with 70% ethanol, a condition known to kill most cells, resulting in a high percentage of PI-labeled cells. In all previous studies where we have used PI staining, this reagent control has verified that the PI staining was working properly.

Immunocytochemistry (ICC)

At the end of the culture period, immunocytochemistry was used to evaluate the extent of proliferation and differentiation of the AHPCs following shockwave exposure (S1 Fig). Briefly, the AHPCs on the coverslips were rinsed with ice-cold 0.1 M PO₄ buffer for one minute and then incubated in 4% paraformaldehyde (PFA) made in 0.1 M PO₄ buffer for 20 minutes. Following fixation, the coverslips were rinsed three times for 7 minutes each with phosphate-buffered saline (PBS) to remove all the fixatives. The coverslips were then incubated in a blocking solution composed of 0.2% Triton X-100 (Thermo Fisher Scientific), 5% normal donkey serum (NDS) (Jackson ImmunoResearch), and 0.4% bovine serum albumin (BSA) (Sigma-Aldrich) in PBS at room temperature for 90 minutes. Primary antibodies—Rabbit Ki-67 (1:300, IgG; Abcam), Mouse TuJ1 (1:200, IgG; R&D Systems), Mouse MAP2ab (1:200, IgG; Sigma-Aldrich), Mouse RIP (1:200, IgG; DSHB, Iowa City, IA), and Mouse GFAP (1:200, IgG; Sigma-Aldrich)—were diluted in the blocking solution. See S1 Table for details on cell markers these primary antibodies are being used to indentify. The cells were incubated in the primary antibody solution overnight at 4° C. The next day, the samples were rinsed four times for 8 minutes each with PBS. To prepare the secondary antibody solution, Donkey Rabbit AF488 (1:300, IgG; Jackson ImmunoResearch), Donkey Mouse AF488 (1:200, IgG; Jackson ImmunoResearch), and Donkey Mouse Cy3 (1:200, IgG; Jackson ImmunoResearch) were diluted using the blocking solution, which also contained DAPI (1:500). The cells were incubated with the secondary antibody solution at room temperature in the dark for 90 minutes and then were rinsed with PBS four times for 8 minutes each. Coverslips were then mounted onto microscope slides with Fluoromount-G with DAPI. Notably, the DAPI counterstaining used throughout the results section was performed on paraformaldehyde fixed cells. The DAPI was not used as a live cell marker. The total number of cells in an image field was determined by counting the number of DAPI stained nuclei.

Leica fluorescent microscope

For cells cultured in 35 mm dishes, the cells were imaged using an inverted Leica fluorescent microscope (Leica DMI4000B; Leica Microsystems) equipped with standard epifluorescence illumination and a Leica DFC310 FX (Leica Microsystems) digital camera. For cells cultured on coverslips, the cells were imaged using an upright Leica fluorescent microscope (Leica DM5000B; Leica Microsystems) equipped with standard epifluorescence illumination and a Q Imaging Retiga 2000R (Q Imaging) digital camera. A 20 objective was used to obtain images for quantitative data analysis, while a 40 objective was used to obtain high magnification images of cellular morphology.

Data acquisition and statistical analysis

The images of the AHPCs were analyzed and quantified using ImageJ software (http://imagej.nih.gov/ij). PI, Ki-67, TuJ1, MAP2ab, and RIP immunoreactive cells were counted using the Cell Counter tool in ImageJ, and the percentage of immunoreactive cells was determined as the number of positively labeled cells divided by the number of DAPI-labeled nuclei. For each independent experiment, ten image fields were acquired for each stain or antibody. Consequently, 20 or 30 image fields per condition were quantified from N = 2 or N = 3 independent experiments, respectively. Then, the percentage of immunoreactive cells was averaged across all these image fields, resulting in one representative measurement per experimental condition. On the other hand, a total of 20 individually indented cells per condition were quantified for the AFM analysis. The control group comprised cells from N = 2 independent experiments, while the shockwave-exposed group comprised cells from N = 3 independent experiments. All means are reported with standard deviation (mean ± SD) or standard error of the mean (mean ± SEM). Graph Pad Prism 10 (Graph Pad Software, Inc., San Diego, CA) was used for ICC/PI, and OriginPro 2022 (64-bit) SR1 version 9.9.0.225 (Academic) was used for AFM statistical analysis and graph-making. Statistical comparisons were conducted on the mean of these replicates (N = 2 or N = 3 replicates per condition), using either an unpaired T-test with Welch’s correction (two-group comparisons) or ordinary one-way ANOVA using Tukey’s multiple comparisons test, with for both ICC/PI and AFM. The significance levels are denoted as follows: * for P < 0.05; ** for P < 0.01; *** for P < 0.001; and **** for P < 0.0001.

The units used for statistical analysis in this study vary depending on the assays and their designs. For example, in the case of the population-averaged parameters (ICC, PI), the unit of analysis is an image field from an independently prepared coverslip, while cells within an image field act as technical sub-samples. In contrast, for ‘single-cell AFM nanoindentation’ experiments, cells are analyzed individually because intercellular mechanical heterogeneity is a well-known factor in this type of measurement [34–36]. In compliance with the standard approach in single-cell AFM experiments [37–39], we therefore used the individual cell, pooled across N = 3 independent preparations, as the statistical unit for the AFM comparisons.

Morphological changes in cells following shockwave exposures

We investigated the morphological changes of AHPC cells subjected to shockwaves with varying intensities and orientations. In control cells (that were not exposed to shockwaves), the actin cytoskeleton was strongly assembled into actin-rich structures with clear assemblies and defined striations contributing to the mechanical strength, and structural integrity of the cell (see Fig 2A,2D). These stress fibers are mainly composed of actin and myosin and are connected to focal adhesion sites on the plasma membrane [40] that ensure mechanical resilience and stable substrate adhesion. Upon shockwave exposure, this cohesive and dense network started becoming weaker and aggregated (Fig 2B-2C) primarily around the cell periphery area. Notably, actin filaments became smoothened and also retracted from the perinuclear area towards the edge (Fig 2E-2F) creating an uneven cytoskeletal distribution. The retracted cells showed a rearrangement of their actin filaments into elongated fibers in the filopodia [41]. Moreover, when shockwaves were delivered from the bottom of the dish, cells displayed membrane blebbing and became more globular and even at forces . Over time, treated cells became more round in shape with surface blebbing but began restoring their shape and cytoskeletal structure around 3 hours post-exposure when they were kept in the incubator, indicating recovery (S8 Fig). In the further study, surface roughness analysis revealed that ‘top-to-bottom’ single shockwaves caused minimal reduction in membrane roughness (RMS: 11.17 ± 1.17 nm) compared to controls (RMS: 11.69 ± 1.69 nm). However, increased shockwave with double magnitude led to a marked decrease in roughness (RMS: 9.21 ± 1.44 nm, in Fig 3A). On the other hand, ‘bottom-to-top’ shockwaves caused further smoothening of cell surface, with roughness values of 9.83 ± 1.10 nm (single) and 8.14 ± 1.08 nm (double exposure), as shown in Fig 3B. These findings suggested that shockwave exposures at higher magnitudes/ intensities and ‘bottom-to-top’ direction weaken the actin cytoskeleton, leading to reduced membrane roughness and altered cell morphology.

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Fig 2. Peak force error images of cells obtained using AFM under different shockwave exposure conditions.

The top section shows images of control cells(A) alongside cells exposed to a shockwave from the top at 14.5 psi(B) and 29 psi(C). The bottom section presents control images(D) along with images of cells exposed to a shockwave from the bottom of the cell dish at 14.5 psi(E) and 29 psi(F). This comparison highlights morphological changes in cell surfaces due to directional shockwave exposure at varying pressures.

https://doi.org/10.1371/journal.pone.0355739.g002

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Fig 3. Effects of shockwave exposure from top-to-bottom (A, C, E, G, I) and bottom-to-top (B, D, F, H, J) directions on the morphological and mechanical properties of AHPCs.

Box plots illustrate comparisons across three conditions: before shockwave (cells were not exposed to shockwaves), after single shockwave, and after double shockwave exposure: under top-to-bottom shockwave exposure for (A) surface roughness, (C) Young’s modulus, (E) deformation, (G) retardation time, and (I) viscosity. Similarly, the impacts of single and double shockwave exposure from the bottom-to-top direction on AHPCs are shown in box plots for (B) surface roughness, (D) Young’s modulus, (F) deformation, (H) retardation time, and (J) viscosity. Significant decreases in roughness, Young’s modulus, viscosity, and retardation time, along with an increase in deformation, are observed after shockwave exposure, particularly following double shockwave exposure in both cases. ‘X——X’ represents the upper and lower extremes of the data, while the error bars extend to a range of 1.5 times the interquartile range (IQR). The mean is depicted by the small box, and the median is indicated by the black line. Statistical significance is denoted as follows: ns (not significant), * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001.

https://doi.org/10.1371/journal.pone.0355739.g003

Influence of shockwave application on cell mechanics

Viscoelastic properties.

Similarly, we investigated the relaxation behavior of AHPCs to understand how different bTBI conditions influence viscoelastic properties that were obtained at varied scan rates and loading velocities. We used ‘Ramp and Hold’ to characterize the cellular surface viscoelasticity by studying their dynamic stress response as a function of applied strain. The AFM tip was approached towards the cell surface at scan rates of 1 Hz, 2.03 Hz, 3.05 Hz, 4.07 Hz, and 5.09 Hz, with corresponding loading velocities of 10 m/s, 20.3 m/s, 30.5 m/s, 40.7 m/s, and 50.9 m/s, respectively. The AFM tip approached the cell during each measurement until a defined loading force threshold was reached. Then, the cantilever base displacement constant was held for 2 seconds before retracting at the same velocity. These time-series force relaxation curves in Fig 4A-4E showed a nonlinear decrease in force after reaching the set point, characteristic of viscoelastic material behavior. At lower scan rates (e.g., 1 Hz), the cell surface showed a slower and more extended force decay since cells had more time to respond to the applied stress. In contrast, at higher scan rates (e.g., 5.09 Hz), the force decayed more steeply and rapidly, indicating a stiffer response due to limited time for viscous deformation due to the fast movement of the probe. However, compared to the controlled AHPCs’ behavior, the relaxation process was similar across all cases (Fig 4A-4E), whether the shockwave was applied from top-to-bottom (Fig 4B-4C) or bottom-to-top (Fig 4D-4E), regardless of the exposure duration of 1.76 ms or consecutive two exposures of 1.76 ms each.

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Fig 4. Representation of force-time curve measured in AHPCs before shockwave exposure (A), after a single shockwave (B, D), and after double shockwave exposures (C, E) from both top-to-bottom (B, C) and bottom-to-top (D, E) directions, with the scan rate ranging from 1 Hz to 5.09 Hz.

The tip approached and detected the cell surface, and during the 2-second holding period, the loading force began to decrease because of the cell’s viscoelastic properties.

https://doi.org/10.1371/journal.pone.0355739.g004

We employed the Kohlrausch-Williams-Watts (KWW) function [33] (see Eq. 5 in section Analysis for Nanomechanical Measurements, an empirical model to describe the dispersion processes in viscoelastic systems to assess the relaxation behavior quantitatively. The relaxation time () for control cells was determined to be . Following a single shockwave exposure (top-to-bottom), the relaxation time showed minimal change at , whereas double shock exposure resulted in a decrease to (see Fig 3G). In contrast, the bottom-to-top shockwave exposure produced more significant effects: single shockwave measurements yielded a relaxation time of , and double shockwave reduced those times even more to 0 (Fig 3H) respectively. The analysis of cellular viscosity also revealed notable trends. Control cells exhibited a viscosity of . After a top-to-bottom single shockwave exposure, the viscosity decreased to , with a further reduction to respectively following double shock exposure (Fig 3I). For cells exposed to bottom-to-top shocks, the viscosity following a single shock was , and this value dropped significantly to respectively after a double shock (Fig 3J). Overall, our results show that although AHPCs’ viscoelastic relaxation behavior was consistent across bTBI conditions, exposure to ‘bottom-to-top’ shockwaves resulted in more noticeable decreases in viscosity and relaxation time, suggesting a higher level of cellular dissipative mechanics impairment.

Assessment of AHPC viability after shockwave exposure

To evaluate whether AHPCs remain sufficiently viable after exposure to shockwaves, we performed viability assessments using propidium iodide (PI) staining at various time points post-exposure. The PI staining performed 24 hours after shockwave exposure showed that exposure to single or double shockwave blast had little to no effect on cell viability shortly after shockwave compared to the control group as the cell viability remained above 99% for all conditions (S2 Fig). However, there was a slight, yet significant, decrease in the percentage of viable cells for the double shockwave exposed samples compared to the single shockwave samples, though both groups remained above 99% (S4 Fig A). Furthermore, cell viability remained high even at the end of the culture period, with the percentage of PI-labeled cells for both shockwave groups at 13 DIV above 99% for all culture conditions (S3 Fig). As with the early time point, there was a slight, yet significant decrease in cell viability for the double shockwave conditions compared to the single shockwave condition, though both groups remained above 98% (S4 Fig B). These results indicated that there was a negligible effect of single or double shockwave exposures from ‘top-to-bottom’ direction on the viability of the AHPCs shortly after exposure or after a longer-term culture.

To reduce the effects of shockwaves traveling through the dish lid, air, and culture media, the shockwave direction was inverted to expose the cells from the ‘bottom-to-top’ direction for a more direct impact. Cell viability was assessed 3 hours after shockwave exposure to evaluate its short-term impacts. Regarding the single shockwave condition, the viability percentages were above 99% for both the control and shockwave-exposed samples (S5 Fig A-C). When comparing the control and double shockwave-exposed samples, the viable cell percentages were both above 98% (S5 Fig D-F). Cell viability was reassessed after 10 DIV to ensure viability at the time of the ICC. Cell viability was high (above 99%) for both the single and double shockwave-exposed AHPCs, confirming the cells’ high viability post-expansion (S6 Fig A-F). With respect to both time periods, results showed that single and double shockwave exposures from the ‘bottom-to-top’ direction had minimal impact on cell survival, with no significant differences in viability compared to their non-exposed controls. Comparing the cell viability results of the single and double shockwave-exposed samples, viability decreased slightly following double shockwave exposure at the 4 DIV, though there were no significant differences between the shockwave conditions at both time periods (S7 Fig A-B). This suggests shockwave exposure at these strengths had little effect on the viability of remaining adherent AHPCs.

Impact of shockwave exposure on AHPC differentiation potential

To evaluate the impact of shockwave exposure on the differentiation and proliferation potential of AHPCs, immunocytochemical analysis was conducted to assess the expression of neuronal, oligodendrocyte, astrocyte, and proliferative markers across all experimental conditions. The results showed there were no significant differences in the expression of any of the antibodies tested between the control and single shockwave exposed samples; however, significant decreases in antibody expression began to be seen following double shockwave exposure (see Fig 5). The percentage of TuJ1-expressing developing neurons was 22.29 ± 15.48% compared to 18.84 ± 14.03% for the control and single shockwave exposure conditions, respectively (Fig 5A-5B’,M). In contrast, there were significant changes following double shockwave exposure as illustrated by the significant decrease in TuJ1-expression, where the percentage decreased from 11.41 ± 1.7% for the control condition to 6.42 ± 0.7% for the double shockwave condition. The percentage of TuJ1 expression following double shockwave exposure was also significantly lower than that seen following single shockwave exposure (Fig 5G-5H’, N, and S4 Fig C). MAP2ab expression showed a minimal decrease following shockwave exposure with expression dropping from 12.54 ± 9.15% to 9.28 ± 11.48% for single shockwave exposure condition and from 11.49 ± 1.3% to 10 ± 1.3% for double shockwave exposure conditions (Fig 5C-5D’, I-J’,M-N). RIP expression slightly decreased, going from 39.4 ± 16.04% for the control condition to 37.29 ± 17.21% for the single shockwave condition (Fig 5E-5F’,M) but showed a significant decrease in expression following double shockwave exposure with the average percentage of RIP-expressing cells dropping from 44.06 ± 3.3% to 31 ± 2.4% (Fig 5K-5L’,N). There was no GFAP expression seen for either the control or shockwave exposed samples. Minimal change was seen in Ki67-expressing proliferating cells, (Fig 5A-5B’,G-H’,M-N). Overall, these results show that single shockwave exposure from ‘top-to-bottom’ direction had little to no effect on cell viability, proliferation, or differentiation while the double shockwave exposure had some significant effect on immature neuron and oligodendrocyte differentiation.

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Fig 5. Differentiation and proliferation of AHPCs at 12-13 DIV following single or double shockwave exposure from top-to-bottom.

Fluorescence images of AHPCs on coverslips at 12 or 13 days in vitro (DIV) following single (A-F’) or double (G-L’) shockwave exposure from above. Cells were single or double immunostained with an immature neuron marker (TuJ1, red; A-B’), a cell proliferation marker (Ki67, green; A-B’), a maturing neuronal marker (MAP2ab, red; C-D’), an oligodendrocyte marker (RIP, red; E-F’), and a cell nuclei marker (DAPI, blue; A-F’). Images A-F were taken using a 20x objective, with a scale bar of 50 µm. Images A’-L’ represent the boxed region in each respective image at a higher magnification, with a scale bar of 25 µm. M: There was no significant difference between the culture conditions. Bars represent the mean percentage of immunolabeled cells, and the error bars represent the standard deviation (± SD). N = 2 independent experiments, with 20 image fields quantified for each condition. N: There was a significant decrease in the expression of TuJ1 and RIP following double shockwave exposure. There was no significant difference between the culture conditions for the remaining antibody labeling. Bars represent the mean percentage of antibody-labeled cells, and the error bars represent the standard error of the mean (±SEM). N = 3 independent experiments; 30 image fields were quantified for each condition.

https://doi.org/10.1371/journal.pone.0355739.g005

However, at 10 DIV following bottom-to-top shockwave exposure, single shockwave exposure showed no consistent trend in antibody expression between control and exposed samples. Double shockwave exposure slightly increased antibody expression, but there were no significant differences between control and shockwave-exposed samples or between shockwave conditions, except TuJ1. TuJ1 expression showed no patterns or significant difference between the controls and single and double shockwave-exposed samples (20.05 ± 1.6% to 18.83 ± 1.8% and 29.12 ± 1.8% to 32.01 ± 2.6%, respectively) (Fig 6A-6B’, G-H’, M, N). Although in comparing single and double shockwave exposures, double shockwave resulted in significantly higher TuJ1 expression (32.01 ± 2.6% versus 18.83 ± 1.8%, respectively) (S7 Fig C). MAP2ab expression demonstrated a consistent, though non-significant, increase from the controls to the shockwave-exposed samples. Expression increased from 12.85 ± 1.1% to 13.79 ± 0.8% following a single shock and 15.4 ± 1.2% to 15.61 ± 1.2% following a double shock (Fig 6C-6D’, I-J’, M, N). No significant difference was found in a comparison of the expression between single and double shockwave-exposed samples (13.79 ± 0.8% and 15.61 ± 1.2%, respectively) (S7 Fig C). Expression of the RIP antibody also demonstrated consistent, though non-significant, increase from the controls to the shockwave-exposed samples. Following single and double shockwave exposures, that expression increased from 21.21 ± 1.19% to 24.17 ± 1.8%, and 20.18 ± 1.3% to 22.52 ± 1.2%, respectively (Fig 6E-6F’, K’-L’, M, N). Between the single and double shockwave exposures, expression decreased from 24.17 ± 1.8% to 22.52 ± 1.2%, which did not indicate a significant difference (S7 Fig C). No GFAP-immunoreactive AHPCs were detected between any of the conditions. Lastly, we selected to use Ki67 immunolabeling with the goal to determine whether cells are generally proliferative versus quiescent following the various shockwave treatments. Since Ki67 is an endogenous nuclear protein expressed during all active phases of the cell cycle (G1, S, G2, and M), while it is absent in quiescent (G0) cells, this approach was useful for estimating the overall growth fraction of the population of cells. Ki67 expression slightly decreased following single shockwave exposure from 3.38 ± 0.7% to 3.25 ± 0.6% but increased following double shockwave exposure from 2.69 ± 0.3% to 3.1 ± 0.3% (Fig 6A-6B’, G-H’, M, N). There were no significant differences between the control samples and shockwave-exposed samples, nor between the single and double shockwave-exposed samples (3.25 ± 0.6% to 3.1 ± 0.3%, respectively) (S7 Fig C).

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Fig 6. Differentiation and Proliferation of AHPCs at 10 DIV Following Single or Double Shockwave Exposure from Bottom-to-Top.

Fluorescence images of AHPCs on coverslips at 10 DIV following single (A-F’) or double (G-L’) shockwave exposure from below. Cells were single or double immunostained with an immature neuron marker (TuJ1, red; A-B’), a cell proliferation marker (Ki67, green; A-B’), a maturing neuronal marker (MAP2ab, red; C-D’), an oligodendrocyte marker (RIP, red; E-F’), and a cell nuclei marker (DAPI, blue; A-F’). Images A-F were taken using a 20x objective, with a scale bar of 50 µm. Images A’-L’ represent the boxed region in each respective image at a higher magnification, with a scale bar of 25 µm. M:There was no significant difference between the culture conditions. N: There was no significant difference between the culture conditions. Bars represent the mean percentage of immunolabeled cells, and the error bars represent the standard error of the mean (±SEM). N = 3 independent experiments; 30 image fields were quantified for each condition.

https://doi.org/10.1371/journal.pone.0355739.g006