Vibration, a treatment for migraine, linked to calpain driven changes in actin cytoskeleton.

Understanding how a human cell reacts to external physical stimuli is essential to understanding why vibration can elicit localized pain reduction. Stimulation of epithelial cells with external vibration forces has been shown to change cell shape, particularly in regards to structures involved in non-muscle cell motility. We hypothesized that epithelial cells respond to vibration transduction by altering proteins involved in remodeling cytoskeleton. Epithelial cells were exposed to vibration and assessed by microscopy, cytoskeletal staining, immunoblotting and quantitative RT-PCR. Here, we report that epithelial cell lines exposed to 15 minutes of vibration retract filopodia and concentrate actin at the periphery of the cell. In particular, we show an increased expression of the calcium-dependent, cysteine protease, calpain. The discovery that cell transitions are induced by limited exposure to natural forces, such as vibration, provides a foundation to explain how vibrational treatment helps migraine patients.

Introduction

People working with vibration intensive machines, like a jack hammer, are exposed to prolonged vibration exposure. This type of stimulus can be deleterious and has been associated with Raynaud’s disease and rheumatoid arthritis [1, 2]. However, brief and localized encounters with vibration is analgesic and is known to reduce pain of such things as injections [3], acute headaches [4] and hair restoration procedures [5]. Clinically, vibration is also used in managing diseases such as meibomian gland dysfunction [6], cerebral palsy [7, 8] and Parkinson’s disease [9].

Although the positive impact of vibration on an organism has been investigated for over a century [10], there is limited information about how the cells that make up the organism respond to vibration. In truth, vibration is thought of as a good way to pop open a cell to get at its intracellular content [11]. However, with the recent re-emergence of mechanobiology [12], there is a new appreciation that natural forces, such as vibration, impact cells in meaningful ways. Therefore, understanding how a human cell reacts to vibration is essential to understanding why vibration can elicit changes in the whole organism.

At a cellular level, external vibration forces have been shown to trigger cell shape changes involving filopodia [13]. Filopodia are antennae-like structures assembled when a cell has the urge to move [14]. Cell motility depends on actin cytoskeleton remodeling [15]. In particular, there is a need to establish filopodia whereby actin associated proteins work in coordination to initiate parallel actin bundles [16-18]. Filopodial linear growth and retraction is the most commonly observed dynamic of filopodia associated with cell movement [19].

In epithelial cells, filopodia are constructed conventionally with actin filament elongation at the tip and parallel actin bundling without actin branching in the shaft [20]. When epithelial cells contact another cell, the cells establish enough receptor-ligand interactions to retract filopodia [21]. Filopodial retraction involves actin depolymerization at the tip and disassembly of actin bundles in the shaft [22]. This type of actin filament remodeling relies on a large collection of proteins [reviewed in [23]] and proteins involved in treadmilling actin [24].

Calpain is one such actin remodeling protein and is required for cell movement [25] and filopodial dynamics [26]. Calpain’s proteolytic activity has been associated with disconnecting integrin from actin filaments [27]. Integrin-containing filopodia are part of establishing mature cellular adhesions [28]. So, calpain’s disruption of focal adhesions would provide an optimal environment for filopodial retraction. In this regard, the overarching hypothesis for this study is that vibration transduction induces calpain-driven cell shape changes.

Materials and methods

Cells

HeLa (ATCC-CCL-165), a human cervical epithelial carcinoma cell line, were grown in Dulbecco’s Modified Eagle Media (DMEM) supplemented with 10% Fetal Bovine Serum and 50 U/ml penicillin, 50 μg/ml streptomycin and 0.5 μg/ml fungizone amphotericin B at 37°C under 95% humidity and 5% CO2. Cultures were plated at 10,000 cells/cm2 two days prior to the experiment and were approximately 75% confluent when vibrational stimulus was applied. This study did not involve human participants. The human cells used in this study are established and commercially available and as such were granted IRB approval to procure and use.

Vibration

Cells were rinsed with and left in PBS (sans Ca2+ and Mg2+) and either placed for 15 minutes on a vibration resistant table [vibration (-)] or exposed to vibration stimulus with an analog vortex mixer (VWR) at 1,200 rpm [vibration (+)]. The use of PBS (-/-) limited potential addition of extracellular calcium and supplement noted in media.

A Teren VM-6370 vibrometer with touch probe was used to measure vibration every minute for 15 minutes. Vibration readings were most consistent when quantified velocity (mm/s) rather than acceleration or displacement, which were variable. Vibration resistant table produced 0 mm/s velocity and vortex set to 1,200 rpm produced 12.09 ± 1.69 mm/s velocity vibration, which was comparable to levels produced by standard equipment in environment (e.g. vibration produced by clock was 3.7 ± 0.1 mm/s and by air handler was 8.3 ± 0.1 mm/s). For comparison, 1,200 rpm was approximately equal to 20 Hz [1 rpm is equal to 0.0167 Hz; https://energyeducation.ca/encyclopedia/RPM], a vibrational stimulus similar to levels used to treat migraines [4].

Chosen vibration stimulus (1,200 rpm) was based on exploratory experiments documenting cell shape of cells left in 37°C CO2 incubator (0.8 ± 0.1 mm/s), exposed to 600 rpm and exposed to 2,700 rpm. Cells left in incubator were similar to vibration (-) cells and cells exposed to 600 rpm had similar, albeit lessened, responses as those seen at 1,200 rpm. Cells exposed to 2,700 rpm (well outside the range of common vibration parameters), detached from the tissue culture vessel. Therefore, subsequent experiments set the vibrational stimulus at 1,200 rpm (12.09 ± 1.69 mm/s velocity).

Microscopy

Micrographs were documented using CellSens imaging software using an inverted, Olympus CKX41 outfitted with phase contrast and fluorescence (excitation 470/40; emission 525/50) using an InfinityHD digital camera.

Cell phenotype

Cells (≥250) from combined field of views were counted as either rounded or not (having extended lamellipodial networks). Individual cell phenotype was quantified using NIH ImageJ software [29] to measure cell area (drawn manually around perimeter including any protrusions) for greater than or equal to 100 randomly chosen cells. Rounded phenotype was characterized by an overall reduction in cell area.

Actin filament staining

Cells were fixed with 3.7% formaldehyde (sans methanol), permeabilized with 0.1% Triton X-100 and stained for actin filaments with 0.165 μM phalloidin-AlexaFluor488 (Molecular Probes). Filopodia length was quantified using NIH Image J software with 400x total magnification field of view calibrated to 450 microns. Filopodia (≥100 with a maximum of five per cell) were delineated manually from cell body to end to measure filopodia length in microns.

Immunodetection

For analysis of calpain-1 protein levels, cell lysates were prepared in lysis buffer (50mM Tris, 1% Triton-X100, 0.1% SDS, 0.5% deoxycholate, 150mM sodium chloride, and 1x protease inhibitor cocktail (Cell Signaling Technology, Danvars, MA). Lysates were cleared by centrifugation and protein concentrations were determined using a protein assay (BioRad) according to manufacturer’s instructions. Samples, 20 μg total protein, were subjected to SDS-polyacrylamide gel electrophoresis on 10% Tris-glycine gels (BioRad) and blotted onto PVDF membranes using Owl semi-dry transfer blot apparatus.

Total protein present in duplicate gel was stained with Acqua Stain protein gel stain (Bulldog Bio) as per manufacturer’s instructions and viewed and analyzed with BioRad Gel Doc EZ Imager Image Lab V3.0 software.

The PVDF membranes were blocked with 5% dry milk in TBS/Tween20 and then membranes were probed with 1) 1:1000 monoclonal primary antibody, mouse anti-human calpain-1 (BioRad, VMA00353) (80kD) or 2) 1:2000 monoclonal primary antibody, mouse anti-human beta-actin control (ThermoFisher, BA3R) (42kD) followed by incubation with horseradish-peroxidase (HRP)-linked goat anti-mouse secondary antibody. Colorimetric HRP substrate was visualized using Opti-4CN detection kit (BioRad) as per manufacturer’s instructions. Membranes were mildly stripped (Pierce, Rockford, IL), of one set of antibodies, tested to ensure detection reagents were removed and then re-probed with the next set of antibodies. The specific band for the protein of interest was identified by its relative electrophoretic mobility (relative front) with respect to the size standard. Density of band was assigned an arbitrary volume. Specific binding was quantified by densitometry using NIH ImageJ imaging software. The protein levels were normalized to internal beta-actin. Immunoblot data represented graphically represents immunoblots performed from different experiments (n = 3).

Calpain inhibition

Cells were pre-treated for 20 hours with DMSO solvent control [inhibitor -] or 100 μM PD150606, a calpain inhibitor (Calbiochem) [inhibitor +]. Although vibration was for only 15 minutes, calpain inhibition is reversible. So, immediately prior to exposing cells to vibration, cells were rinsed with PBS and treated with a fresh dose of DMSO or calpain inhibitor in PBS. Individual cell phenotype was quantified by cell area as described above.

Quantitative Real-Time Reverse-Transcription Polymerase Chain Reaction (qRT-PCR)

SurePrep™ TrueTotal™ RNA Purification Kit (Fisher) was used to extract total RNA as per manufacturer’s instructions. RNA quantification at A260 used to standardize amount RNA (10 ng/μL) loaded into High-Capacity RNA to cDNA kit cDNA for reverse transcription. One-tenth of the cDNA was subjected to qRT-PCR using TaqMan® Fast Advance Master Mix (Applied Biosystems) in conjunction with primer probes (ThermoFisher): human calpain-1 (Hs00559804_m1), E-cadherin (Hs01013958_m1) and GAPDH (Hs0392907_g1) in conjunction with a QuantStudio 5 Real-Time PCR instrument. Melt curves indicate the primer set produced one product. The ΔΔCt method calculation was used for comparing expression levels between cells exposed to vibrational stimulus or not.

Statistical analysis

Two-tailed T-test (Two-sample assuming equal variances) for analysis of cell area and filopodial length, which had only two groups and ANOVA supplemented with Tukey HSD analysis was performed with Excel data analysis package.

Results

Cells exposed to vibration were more susceptible to losing prominent filopodia

Cells growing in non-confluent adherent conditions had prominent filopodia and lamellipodia, a characteristic of an established cell culture (. After exposure to 15 minutes of vibrational stimulus, a significant percentage of cells retracted these actin-rich plasma membrane protrusions and displayed a round shape indicative of a less adherent cell (. The number of cells displaying each cell shape was counted and calculated as a percentage of cells that were ‘round’[round phenotype in 9.0 ± 0.5% cells without vibration vs 57.6 ± 10.3% cells with vibration].

Vibration induces a cell shape change consistent with loss of established filopodia.

Representative phase contrast micrographs of HeLa cells without vibration (A) and with vibration (B) at 200x total magnification. (C) The perimeter of each cell (≥100) was delineated including any protrusions and the cell area was measured (arbitrary units, AU). Data shown as average cell area (AU) for all cells measured (±sem for three independent experiments). * = p < 0.05 Vibration (+) [solid bar] vs. Vibration (-) [striped bar].

Retraction of protrusions displayed by the round shaped cells resulted in a significant reduction in the average cell area of cells exposed to vibration (. Thereby, cell area was used subsequently as a quantifiable criteria for assessing round shape stimulated by vibration.

Cells exposed to vibration have altered actin filament assembly

Filopodia are exploratory structures constructed from actin filaments [reviewed in [30]]. The retraction of filopodia indicated that vibration was impacting efficient actin filament polymerization and bundling, steps in filopodia dynamics [reviewed in [31]]. Cytoskeletal architecture was assessed with phalloidin staining of actin filaments, whereby straight needle-like rays indicate filaments bundled in a filopodia and clouds of homogenous intensity around a bright point indicate short filaments [32]. Cells growing with no vibration displayed rays indicative of filopodia (, while cells exposed to a vibrational stimulus displayed clouds of actin staining around the cell periphery indicative of shortened filaments (. Stress fibers are distinct from filopodia in that they traverse the entirety of the cell.

Vibration alters actin filament organization.

Representative fluorescence micrographs of HeLa cells without vibration (A) and with vibration (B) stained for actin filaments. Filopodial length calculated using ImageJ image analysis software. Field of view diameter for 400x total magnification set to 450 microns. Individual filopodia (≥100, with maximum of five per cell) were marked from cell body to end for length measurement. Stress fibers running across entire cell body were excluded. Data shown as average filopodial length in microns (±sem for three independent experiments) (C). * = p < 0.05 Vibration (+) [solid bar] vs. Vibration (-) [striped bar].

It was presumed that as actin bundles destabilize and filaments shortened, phalloidin staining density, quantified by image analysis, would increase. Cells exposed to vibrational stimulus had a decrease in average filopodial length in comparison to cells not exposed to vibration (. The cell staining pattern indicated a redistribution of bundled actin filaments to unbundled, shortened filaments, particularly at the cellular periphery. This pattern of staining is indicative of actin filament detachment from integrin or actin anchoring proteins.

Vibration altered intracellular levels of protein

The results from the actin filament experiments indicated that vibration induced actin unbundling, which is mediated by actin-binding proteins and their associated proteases. To investigate whether actin associated proteins were involved, gel electrophoresis was performed on cell lysates from HeLa exposed or not exposed to vibrational stimulus (Fig 3A). An assessment of total protein relative volume (AU) revealed cells exposed to vibration had significant increases in intracellular levels of several proteins, whose band size was in the vicinity of the 72 kD protein marker (. Assessment of other bands, which showed similar or coincident decrease in protein levels (e.g. bands 1 and 6), mitigated the possibility that these alterations in proteins levels were simply a product of unequal loading of protein amounts between the samples.

Fig 3

Vibration altered intracellular protein content.

Representative protein bands obtained from HeLa cells without vibration and with vibration. Cells were lysed for total protein, separated by vertical electrophoresis and stained for total protein. Marker indicates 10 molecular marker bands corresponding to protein size (A). Seventeen bands from each lane were aligned based on relative front in comparison to relative front of marker bands. Arrow indicates placement of the 72kD marker. An arbitrary volume unit was assigned to each band by analysis software (B). Larger volumes were assigned as band density increased indicating an increase in protein levels. Band volumes for cells exposed to vibration are displayed as relative volume difference to cells not exposed to vibration. Numbers great than one indicate increase in vibration stimulated cells. Data shown as average relative difference volume (±sem for three independent experiments). Of note are bands 8, 9 and 10 (clustered around 72kD) that were increased in HeLa with vibration in comparison to HeLa without vibration. Alternatively, bands 1and 6 were decreased in HeLa with vibration in comparison to HeLa without vibration.

Vibration increased the intracellular level of calpain protein

A literature search on actin associated proteins with a relevant size led to calpain-1, a ubiquitous Ca2+-activated protease. First, calpain inhibition leads to a decrease in cell detachment rates [25], which infers calpain involvement in increasing cell detachment or cell rounding. Second, calpain cleaves actin-associated cytoskeletal proteins [33], which infers calpain involvement in actin unbundling. Lastly, calpain-1 is 80 kD in size [34].

As determined by immunoblotting, calpain-1 levels increased as HeLa were treated with increasing vibrational stimuli. Beta-actin was detected at similar levels in each sample indicating that calpain increase was not a consequence of loading differences (. Relative calpain protein was significantly increased in cell stimulated with vibration representing 1200 rpm (V++) (. Thereby, a vibration stimulus increased calpain-1 (also known as μ-calpain) protein levels.

Vibration stimulates increase in calpain protein.

Representative immunoblot of cytoplasmic lysates from HeLa cells without vibration (vibration -) or with increasing levels of vibration [vibration (+) and vibration (++) representing 600 and 1200 rpm, respectively]. Lysates probed for calpain-1 [top panel] and beta-actin [bottom panel] as a loading control. Similar levels of beta-actin support that calpain increase was not a consequence of loading differences. Marker indicates molecular marker bands corresponding to protein size (kD) (A). Relative calpain-1 protein, a ratio of the calpain-1 to beta-actin band peak densities for each immunoblot (n = 3), expressed as average (± sem for three independent experiments). (B). * = p < 0.05 Vibration (++) [dark, solid bar] vs. Vibration (-) [striped bar]. No significant difference noted between Vibration (+) [gray bar] vs. Vibration (-).

Calpain inhibition attenuated vibration-induced cell shape change

Immunoblotting provides support about increases in calpain protein levels. However, it does not address calpain activity. Is calpain actively involved in cell rounding in response to a vibrational stimulus? In this regard, HeLa were pre-treated and exposed to vibration in the presence of a calpain inhibitor.

As previously documented, after exposure to 15 minutes of vibrational stimulus, a significant percentage of cells displayed a reduced cell area in comparison to cells not exposed to vibration. [note: the data presented here is independent of the data presented in previous figures]. In the presence of calpain-inhibitor a significant percentage of cells were unable to take on the rounded phenotype and as such did not display the reduced cell area displayed by their counterparts that were exposed to vibration without calpain inhibitor (. Therefore, vibration-stimulated changes in cell shape involve a pathway in which calpain is actively catalyzing the degradation of one of its substrates.

Inhibition of calpain activity attenuates vibration induced cell shape change.

HeLa cells were pre-treated overnight with DMSO (solvent control) [calpain inhibitor (-)] or with calpain inhibitor, PD150606 [calpain inhibitor (+)]. HeLa cells were washed and re-treated immediately prior to treated without vibration (-) [striped bar] or with vibration (+) [solid bar]. The perimeter of each cell (≥100) was delineated including any protrusions and the cell area was measured (AU). Data shown as average cell area (AU) for all cells measured (±sem for three independent experiments). * = p < 0.05 Vibration (+)/Calpain(-) [solid bar] vs. Vibration (-)/Calpain(-) [striped bar]. ** = p < 0.05 Vibration (+)/Calpain Inhibitor (+) vs. Vibration (+)/Calpain Inhibitor (-). No significant difference was noted between Vibration (+)/Calpain Inhibitor (+) and other sample groups [Vibration (-)/Calpain Inhibitor (+) or Vibration (-) / Calpain Inhibitor (-)].

Vibration altered calpain gene expression

As was noted, calpain is a ubiquitously expressed protein. It was presumed that calpain involvement in vibrational-induced cell shape changes was due to resident calpain and not as a consequence of enhanced calpain gene expression. Current trends suggest that an increase in protein levels does not necessitate an increase in gene expression [35]. However, qRT-PCR utilizing calpain-primers detected a significant fold increase in calpain expression ( indicating that vibration enhances calpain gene expression.

Vibration alters calpain gene expression.

RNA isolated from HeLa cells without vibration or with vibration was subjected to reverse transcription. Resultant cDNA was used as a template for qPCR using primers for calpain and E-cadherin, an epithelial marker. Threshold cycles were normalized to GAPDH, a loading control. A fold difference of expression comparing vibration to no vibration of HeLa was calculated with the ΔΔCT method where no change is indicated by “1”. No reverse transcriptase and no template controls showed no product.

To mitigate the possibility that vibration was inducing gene expression across the genome (rather than specifically at calpain), qRT-PCR was run on additional relevant proteins. E-cadherin was chosen because enhanced calpain expression had been associated to decreased E-cadherin expression [36]. As expected, E-cadherin expression was down-regulated indicating that the enhanced calpain expression was specific to calpain.

Discussion

The data presented here shows that epithelial cells change their shape in response to vibration. In particular, vibration stimulates a rounded shape associated with reduced cell area, actin reorganization leading to reduced filopodial length and an increase in expression of the calcium-dependent, cysteine protease, calpain-1 (μ-calpain).

Sub-confluent cultures and scratch wound assays (a portion of a confluent monolayer is removed) establish a cellular growth environment where cells crawl to find other cells. This urge to move leads to cell shape changes associated with remodeling of actin cytoskeleton [15] and is directed by antennae-like structures called filopodia [14]. The filopodia-making machine uses a convergent elongation mechanism, where filaments undergoing persistent elongation at the barbed-end eventually bundle to form the filopodia [16-18]. Retraction or loss of filopodia depends on destabilizing the barbed-end near the plasma membrane and deconstructing the filament bundles along the shaft.

Based on current findings, sub-confluent cervical epithelial cells (HeLa) cell monolayers that were exposed to vibration displayed a loss or retraction of filopodia. Vibration consistently led to cells that displayed a rounded phenotype with reduced cell area. Actin filament staining showed concentrated plaques of actin filaments around the periphery of the cell. While the remaining cells (not rounded) were observed to have a lamellipodial network consistent with shorter, branched filaments [37], which was associated with a reduction in filopodial length.

The intensity of the actin filament staining in the rounded cells was striking and was reminiscent of cells treated with cytochalasin, which disrupts actin assembly [38]. It appeared as if the filaments had collapsed or folded onto the cell, similar to plectoneme formation during nucleic acid supercoiling. This phenomenon was previously described as a mechanism to create pools of actin that could be recruited into lamellipodia [39]. The actin pools indicate depolymerization, an increase in capping and branching of the filament or a detachment from the membrane.

Congruent to the findings presented here, calpain is known to target actin membrane anchoring proteins such as vinculin in epithelial cells [40] and ezrin in platelets [41]. The data presented here shows that a 15-minute vibration exposure leads epithelial cells to increase expression of calpain-1. Furthermore, calpain activity drives vibration-induced cell rounding and associated reduction in cell area. This data together with other findings make membrane detachment of filopodial filament a feasible mechanism for how vibration induces filopodial retraction. Furthermore, observed increases in calpain expression, which could account for the significant percent change [2.7% ± 1.3] in calpain protein levels in cells with vibration vs. cells without vibration, are supported in whole animal studies where calpain mRNA increases were found in rabbit muscle tissue 45 days after exposure to hind leg vibration [42].

In this regard, calpain inhibition is expected to stabilize filopodia leading them to maintain focal contact with the plasma membrane through to the extracellular matrix at their tips. When epithelial cells were exposed to vibration in the presence of a calpain inhibitor, filopodial structures remained, which reestablished cell area similar to cells not exposed to vibration. However, others have shown in fibroblast cells that calpain inhibition actually prevented focal contact formation [33]. This paradox in reported effects of calpain inhibition has been noted previously in the literature [26] and assumedly hinges on the cell type utilized in various studies. Fibroblast-type cells, such as NIH3T3, behaving differently from epithelial cells, such as HeLa.

Clinically, vibration (kinetic oscillation) is used as an analgesic for migraine pain [4, 43]. However, there is a paucity of information to explain how vibration works at a cellular level. Others have shown that vibration reduces calcitonin-gene related peptides (CGRP) [44]. CGRP are involved in proinflammatory responses noted in migraine pathophysiology [45] and whose reduction has been shown to alleviate migraine pain in animal models [46]. CGRP increases import of calcium into cells [47, 48] indicating that reduction of CGRP would likely lead to reduction in calcium import. The data presented here indicates that vibration leads to an increase in calpain-1 levels. Calpain-1 activity is reliant on micromolar levels of intracellular calcium [49].

Together these findings imply that vibrational analgesia is a consequence of calcium signaling. This idea is supported by studies showing that mechanical stimulus leads to a significant cellular calcium response coordinating calcium influx into the cell and calcium release from the ER [50] As reported previously [51], the calpain inhibitor reported here, PD150606, works by blocking calpain’s calcium binding site. As noted, this calpain inhibitor abrogated the impact of vibration lending further support to vibration mechanotransduction being driven by intracellular calcium levels. In future works, it would be interesting to perform cell fate mapping by pulsing cells with calcium to control for actin filament retraction.

The molecular processes described, such as the induction of a rounded cell shape, in conjunction with decreased expression of E-cadherin, an epithelial gene, suggests that vibration prompts the first steps of an epithelial-mesenchymal transition (EMT) [52]. Calcium signaling has also been shown to induce EMT [53]. That being said, the concept that epithelial cell transition can be mediated by cellular exposure to natural forces, like vibration, is intriguing. Moreover, the reversible nature of EMT on the onset [54] may explain why brief, localized exposure to vibration can be beneficial to human health, while prolonged exposure can be deleterious.

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Reviewer #1: 1.- In figure 2 it is suggested to count the filopodia per cell instead of quantifying the intensity of phaloidin-488, since in the images shown by the authors, these structures are easily recognizable and quantifiable. If possible, measure the length of the filipodia in both conditions.

2.- In figure 3, it is strongly suggested to show the protein gel pattern together with the densitometry.

3.- In figure 4, how many technical replicates were made? It is necessary to do a densitometry analysis of at least two technical replicates to compare between conditions. Also, the image has no captions to recognize what conditions each line has.

4.- In all the experiments the author did not mention how many technical replicates were carried out in each experiment. This information is neccesary.

5.- In figure 1 and 5, the image shows cells with a rounded phenotype. Since this variable is presumably not qualitative, the t-student test is not adequate. Please describe in more detail how this analysis was performed or a chi-square test is suggested.

Reviewer #2: This study analyzes the impact of vibration on filopodia retraction and calpain levels in epithelial cell lines, finding that this type of stimulus induces calpain-driven changes in cell shape.

Raynaund’s disease and rheumatoid arthritis have been associated with jobs that involve the exposure to intense vibration. While, clinical vibration helps in the managing of some diseases (Meibomian gland dysfunction and Parkinson’s), and the brief and localized vibration in the analgesic effect and the reduction of some pain as acute headaches. Thus, prolonged exposure to vibration can be deleterious, while a localized exposure can be beneficial to human health. Therefore, understanding how a human cell reacts to vibration is essential to understanding why vibration can elicit changes in the whole organism either with a positive or negative effect. Then, this is an interesting study because can provide useful information about the cellular transition induced by vibration and contribute to understanding how vibrational treatments could help for pain reduction.

Specific comments:

1) The main concern is that some of the quantitative results are based on subjective observations. The percentage of cells with a rounded phenotype was determined in Figure 1. However, the shape changes are heterogeneous and the rounded phenotype criteria are not specified, such as measuring the cell area that could provide more accurate and measurable results.

2) Figure 2 shows a slight but significant difference in fluorescence intensity per cell area after vibration treatment. Changes in shape would imply a reorganization of actin cytoskeleton and not necessarily an increase in the number of actin filaments, since a greater complexity of the filaments is not observed. In this sense, it is kindly suggested that the exposure times of the captured images should be specified, since automatic image acquisition may result in different capture times and consequently give errors in fluorescence intensity measurements. If the above-mentioned problem did occur, correct accordingly.

3) The authors state that vibration altered intracellular levels of some proteins. Figure 3 shows the result of one experiment since no standard deviations are shown. A single experiment and without showing loading controls are not enough for such assertion. Therefore, the analysis of the densitometry average of at least three replicates and statistical analysis is necessary. Besides, it is kindly suggested to show the graph in a single plane to facilitate its interpretation.

4) Figure 4 is a representative immunoblot, therefore for an accurate quantification it is necessary to perform and plot the densitometry of the bands of calpain (sample) and beta-actin (loading control) of at least three replicates, normalize the results with respect to the loading control and perform the corresponding statistical analysis.

5) As in Figure 1, in Figure 5 it is necessary to define the criteria to consider when a cell is rounded, considering the heterogeneity of shapes observed. In support of this observation, in Figure 1 the percentage of cells without vibration treatment is about 7.5%, while in Figure 5 the percentage of cells in the same conditions (without exposure to vibration and without the calpain inhibitor) is about 40%. Similarly, in Figure 1 the percentage of cells treated with vibration is about 50%, while in Figure 5 the percentage of cells in the same conditions (with exposure to vibration and without the calpain inhibitor) is 75%. These discrepancies may be the result of the use of subjective criteria. Therefore, as in Figure 1 measuring cell area and set specific ranges for extended vs. rounded cell may be useful.

6) Finally, Figure 6 shows that the vibration increase calpain transcript levels (close to fivefold) suggesting an up-regulation of its gene expression. As requested above, a quantitative analysis of the results in Figure 4 and their comparison with the results in Figure 6 would provide clues whether the regulation of the calpain gene is at the transcriptional and/or translational level.

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Reviewer #1: No

Reviewer #2: No

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2 Mar 2022

February 17, 2022RE: Response to Reviewers concerning PONE-D-21-39301

We greatly appreciate the opportunity to revise our manuscript PONE-D-21-39301 entitled “Vibration, a treatment for migraine, linked to calpain driven changes in actin cytoskeleton”.

All journal requirements have been addressed.

1) We have ensured that the manuscript has met style requirements based on provided links, particularly appropriately formatted heading and figure labels and removing funding information from acknowledgements. 2) We have matched and provided grant numbers for the award received from Carver. 3) We have removed the four mentioned ‘data not shown’ references from the manuscript as the comments did not enhance the progression of ideas being presented by the study. 4) We have included a full ethics statement in the “Methods” section of the manuscript. 5) We have provided all original, uncropped and unadjusted images mentioned in the manuscript in a public data repository that can be found at the open science framework (OSF) at the following link. https://osf.io/t64jg/?view_only=7f2c9fbd69004b4782aea68aa86fe736

Response to Reviewer’s Comments

Based on the editor’s comments, we have responded to both reviewer’s recommendations as follows.

Reviewer #1

Overall, we greatly appreciate the comments made by reviewer #1 as they have enhanced the statistical soundness and data availability of our study. As an aside, this is our first experience publishing in an open access journal and we were previously unaware of public data repositories. In this regard, we have created an open access link to our data at OSF so that all data is electronically available.

Specifically, all comments made by reviewer #1 have been addressed.

1.- In regards to the data concerning vibrational impact on actin filament assembly, particularly filopodia (figure 2), we have embraced the reviewer’s comment that these structures are easily recognizable and quantifiable. Therefore, we replaced the staining intensity data with data where we measured the length of the filipodia (as suggested). As such, we replaced figure 2C with the updated figure.

2.- In regards to the data concerning vibration altering intracellular levels of protein (figure 3), we have added figure 3A, which shows the protein gel pattern in addition to the densitometry provided in the original version.

3.- In regards to the data concerning vibration increasing intracellular level of calpain protein (figure 4), we have added figure 4B, which is a densitometry analysis for three technical replicates. We are uncertain why the image had no captions as the figure submitted had these captions. However, perhaps now that the figure has been revised to include figure 4B, these captions are evident. In addition, reference to captions are within the figure legend. Additionally, all original, uncropped and unadjusted pictures of gels with annotations are provided in the public repository.

4.- We have fixed our oversight in regards to being explicit about experimental replication. The materials and methods and figure legends now reflect how many technical replicates were performed. Additionally, data from each replicate is provided in the public repository.

5.- In regards to the statistical analysis performed on the rounded phenotype data presented in figures 1 and 5, we have taken the reviewers suggestion and enhanced the statistical analysis of this data. For the revision, we have used an Excel data analysis package with XReal stats, to perform a one-factor ANOVA supplemented with a Tukey HSD post-hoc analysis within and between groups. The method for this analysis with the addition of technical replicates was updated in the materials and methods. As an aside, based on reviewer #2 comments, the ‘rounded phenotype’ counts were replaced with cell area.

We have revised the manuscript to include all of reviewer #1 comments and appreciate the comments which we believe have enhanced the statistical robustness and data availability of the manuscript.

Reviewer #2

Overall, we greatly appreciate reviewer #2’s comments that this study is of interest and could make a worthy contribution to understanding vibrational treatments. The comments concerning how to improve our subjective observations has substantially improved the quality of this work and we value the reviewer’s insight.

Specifically, all comments made by reviewer #2 have been addressed.

1) In regards to our observation that cells exposed to vibration were more susceptible to losing prominent filopodia (Figure 1), we initially assessed this phenomenon using a rounded phenotype. However, reviewer #2’s insight that cell area would be more quantifiable and less subjective was incredibly valuable. As such, we reassessed the data using cell area and have replaced the original figure 1C showing rounded phenotype with cell area data (revised Figure 1C). In addition, we have added a section “cell phenotype” to the materials and methods to clearly provide the criteria used to assess the ‘round phenotype’. We believe that this has provided more accurate and measurable results.

2) In regards to the data concerning vibrational impact on actin filament assembly, particularly filopodia (figure 2), we noted the reviewer’s comment that the fluorescence intensity per cell area would depend on image acquisition and implies an increase in the number of actin filaments. As we did not suppose that there was an increase of actin filaments, but rather a rearrangement of actin filaments corresponding with a retraction of filopodia, we understand the reviewer’s concerns. As such and based on a suggestion made by reviewer #1 that filopodia were easily recognizable, we have reassessed our data set by measuring filopodial length rather than fluorescence intensity. Therefore, we replaced the staining intensity data with data where we measured the length of the filipodia (replaced figure 2C with the updated figure). In addition, we added a section to the materials and methods to describe this process. We believe this has resolved the technical errors that may have accompanied our original assessment of the data. In addition, the statistical significance of the data was enhanced.

3) In regards to the statistical robustness of the data concerning vibration altering intracellular levels of protein (figure 3), we have added a representative picture of one of three gels (Figure 3A) and provided the average and error for three independent experiments (revised Figure 3 to figure 3B including error bars). The 2-D presentation of the graph was of concern because it impeded interpretation of the data. We realized that interpretation of this data would make it necessary to assess the vibration (+) band volumes in relation to the vibration (-) ones. Therefore, we graphed the data in one dimension utilizing the relative difference in volumes between vibration (+) and vibration (-). In addition, so that the gel picture and graph aligned, we converted the 2-D column graph to a 1-D bar graph. We agree with reviewer #2 that the single plane in addition to the incorporation of error bars facilitates the interpretation of data presented about intracellular levels of protein.

4) In regards to the data concerning vibration increasing intracellular level of calpain protein (figure 4), we have added figure 4B, which is a densitometric analysis of calpain levels in relation to loading controls and includes data from three independent experiments. The corresponding statistical analysis was provided in the figure legend and marked on the figure. Additionally, all original, uncropped and unadjusted pictures of the gels with annotations are provided in the public repository.

5) As in Figure 1, we have reassessed data concerning the ‘rounded phenotype’ in the absence or presence of calpain inhibitor (figure 5) utilizing the cell area criteria defined in this revision of the manuscript. The cell area criteria suggested by reviewer #2 is indeed more robust quantitatively and less subjective, as noted by the attenuated discrepancies between figure 1 and figure 5 data. As noted by the reviewer data from the original submission using the ‘rounded phenotype’ had vibration (-) cells at 7.5% rounded in figure 1 and 40% rounded in figure 5. Utilizing the cell area criteria vibration (-) cells had a 20.3 ± 1.6 area in figure 1 and 31.5 ± 12.1 area in figure 5. Additionally, data from the original submission using the ‘rounded phenotype’ had vibration (+) cells at 50% rounded in figure 1 and 75% rounded in figure 5. Utilizing the cell area criteria vibration (+) cells had a 10.8 ± 1.8 area in figure 1 and 9.1 ± 3.1 area in figure 5. As suggested by reviewer #2, we believe that the discrepancies noted in the original submission were a result of the use of a subjective criterion and that measuring cell area was useful to address these discrepancies. As such, Figure 1C and figure 5 from the original submission was replaced with Figure 1C and figure 5 in the revision which showed data using cell area.

6) In regards to data concerning vibration altering calpain gene expression, reviewer #2’s suggestion to analyze protein levels (figure 4) allowed us to compare how the increase of calpain transcript levels corresponded to an increase in protein levels. We determine that the change in protein levels was 2.7 ± 1.3 aligned well with the 4.8 ± 0.7 fold increase noted in calpain transcripts. We have added a comment to the discussion about this point.

In summation, we found great value in the reviewer’s comments. We believe that their expertise and insight has allowed us to revise the manuscript in ways that have transformed the study from being descriptive to being highly quantitative with objective criteria and robust statistical significance.

We greatly appreciate the reviewers comments that our work is relevant and an interesting topic of study because we are earnest in our belief that the study is of import.

Sincerely,

Adriana J. LaGier, Ph.D.

Associate Professor of Biology

alagier@grandview.edu

515-263-2874

Submitted filename: Response to Reviewers.docx

Click here for additional data file.

30 Mar 2022

31 Mar 2022

Journal request has been addressed. We have reviewed our reference list to ensure that it is complete and correct. We have not retracted any cited papers. However, we found two typographical errors that we have fixed (in reference number 42 the journal title listed twice and in reference number 44 the authors names were all capitals).

All Reviewers comments have been addressed. We have also addressed and revised the article based on all suggestions made by both reviewers.

In response to reviewer #1, we used a T-test, instead of an ANOVA, to analyze data presented in figures 1 and 2. As the result was the same, there were no changes made to the figures. However, we added this analysis to the materials and methods. We also updated the datasets with the new analysis at our public data repository. The new analysis (documents S1.4 Dataset Cell Shape Area Ttest and S2.5 Dataset Cell Actin Filament Ttest) can be found at the open science framework (OSF) at the following link. https://osf.io/t64jg/?view_only=7f2c9fbd69004b4782aea68aa86fe736

In response to reviewer #2, we fixed all the typographical errors mentioned and fixed two additional spacing errors.

Submitted filename: Response to Reviewers minor.pdf

Click here for additional data file.

5 Apr 2022

Vibration, a treatment for migraine, linked to calpain driven changes in actin cytoskeleton

PONE-D-21-39301R2

Dear Dr. Adriana J. Lagier,

We’re pleased to inform you that your manuscript has been judged scientifically suitable for publication and will be formally accepted for publication once it meets all outstanding technical requirements.

Within one week, you’ll receive an e-mail detailing the required amendments. When these have been addressed, you’ll receive a formal acceptance letter and your manuscript will be scheduled for publication.

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Kind regards,

Patricia Talamas-Rohana, Ph.D.

Academic Editor

PLOS ONE

Additional Editor Comments (optional):

Reviewers' comments:

Reviewer's Responses to Questions

Comments to the Author

Reviewer #1: All comments have been addressed

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2. Is the manuscript technically sound, and do the data support the conclusions?

The manuscript must describe a technically sound piece of scientific research with data that supports the conclusions. Experiments must have been conducted rigorously, with appropriate controls, replication, and sample sizes. The conclusions must be drawn appropriately based on the data presented.

Reviewer #1: Yes

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3. Has the statistical analysis been performed appropriately and rigorously?

Reviewer #1: Yes

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4. Have the authors made all data underlying the findings in their manuscript fully available?

The PLOS Data policy requires authors to make all data underlying the findings described in their manuscript fully available without restriction, with rare exception (please refer to the Data Availability Statement in the manuscript PDF file). The data should be provided as part of the manuscript or its supporting information, or deposited to a public repository. For example, in addition to summary statistics, the data points behind means, medians and variance measures should be available. If there are restrictions on publicly sharing data—e.g. participant privacy or use of data from a third party—those must be specified.

Reviewer #1: Yes

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5. Is the manuscript presented in an intelligible fashion and written in standard English?

PLOS ONE does not copyedit accepted manuscripts, so the language in submitted articles must be clear, correct, and unambiguous. Any typographical or grammatical errors should be corrected at revision, so please note any specific errors here.

Reviewer #1: Yes

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6. Review Comments to the Author

Please use the space provided to explain your answers to the questions above. You may also include additional comments for the author, including concerns about dual publication, research ethics, or publication ethics. (Please upload your review as an attachment if it exceeds 20,000 characters)

Reviewer #1: (No Response)

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Reviewer #1: No

11 Apr 2022

PONE-D-21-39301R2

Vibration, a treatment for migraine, linked to calpain driven changes in actin cytoskeleton

Dear Dr. LAGIER:

I'm pleased to inform you that your manuscript has been deemed suitable for publication in PLOS ONE. Congratulations! Your manuscript is now with our production department.

If your institution or institutions have a press office, please let them know about your upcoming paper now to help maximize its impact. If they'll be preparing press materials, please inform our press team within the next 48 hours. Your manuscript will remain under strict press embargo until 2 pm Eastern Time on the date of publication. For more information please contact onepress@plos.org.

If we can help with anything else, please email us at plosone@plos.org.

Thank you for submitting your work to PLOS ONE and supporting open access.

Kind regards,

PLOS ONE Editorial Office Staff

on behalf of

Dr. Patricia Talamas-Rohana

Academic Editor

PLOS ONE