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Article

Quantum Dots Affect Actin Cytoskeleton Reorganization, Resulting in Impaired HeLa and THLE-2 Cell Motility

by
Mileah Metcalf
,
Abhishu Chand
and
Kyoungtae Kim
*
Department of Biology, Missouri State University, 901 S National, Springfield, MO 65897, USA
*
Author to whom correspondence should be addressed.
Micro 2025, 5(2), 29; https://doi.org/10.3390/micro5020029
Submission received: 10 April 2025 / Revised: 28 May 2025 / Accepted: 4 June 2025 / Published: 12 June 2025
(This article belongs to the Section Microscale Biology and Medicines)
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Abstract

:
Quantum dots (QDs) are nanoparticles with intrinsic fluorescence. Recent studies have found that metal-based QDs often impart toxic effects on the biological systems they interact with. Their undefined limitations have offset their potential for biomedical application. Our study aimed to address the research gap regarding QDs’ impacts on the intracellular actin cytoskeleton and the associated structures. Our XTT viability assays revealed that QDs only reduced viability in transformed human liver epithelial (THLE-2) cells, whereas HeLa cells remained viable after QD treatment. We also used confocal microscopy to evaluate the morphological changes in THLE-2 induced by QDs. We further investigated cell protrusion morphology using phalloidin-Alexa488 which selectively labels F-actin. The fluorescent microscopy of this phalloidin label revealed that QD treatment resulted in the redistribution of actin filaments within both THLE-2 and HeLa cells. We also report that the average number of focal adhesions decreased in QD-treated cells. As actin filaments at the cell are peripherally linked to the extracellular matrix via talin and integrin and are thus a crucial component of cell motility, we conducted a migration assay. The migration assay revealed that cell motility was significantly reduced in both THLE-2 and HeLa cells following QD treatment. Our findings establish that the internalization of QDs reduces cell motility by rearranging actin filaments.

1. Introduction

Within the last ten years, interest in quantum dots (QDs) has steadily surged. QDs are a popular subcategory of nanomaterials, ranging from 2 to 10 nm. Conjugating specific ligands to the exterior of QDs drastically alters the way they interact with their surrounding environment. Many studies have conjugated QDs to chemical ligands like folate or folic acid [1,2,3,4,5,6,7] or biomolecule ligands like amino acids [8,9,10]. These specific ligands allow for the cell-specific internalization of QDs. The ability of QDs to also be conjugated to chemotherapeutic agents like gefitinib and doxorubicin further elevates their potential within the biotech industry [11,12,13]. In QDs made from heavy metals, particularly those in the periodic table groups II–VI, their UV fluorescence is brighter and more stable. QDs composed of CdSe or CdTe cores are incredibly common as the quantum yield provided by these cores is resistant to degradation and oxidation [14].
Recently, research has indicated that the functionality of heavy metal QDs may be limited due to their inherent toxicity. Heavy emphasis has been placed on cadmium QDs due to their commonality and well-documented toxic effects [15,16,17,18]. To mitigate the toxicity of the inner cadmium core, a shell may be synthesized around QDs. ZnS is one common example shown to reduce the cadmium core’s toxicity in some cases [19,20,21]. The ZnS shell may not fully prevent the heavy metal core’s toxicity as toxicity has been confirmed in many cancerous cell lines like HeLa and HepG2 despite the presence of the ZnS shell surrounding the CdSe QDs [22,23,24,25,26]. Recently, non-cancerous liver cell line THLE-2 was identified to be highly sensitive to the toxic effects caused by CdSe/ZnS QDs [27]. To corroborate the loss in viability and the generation of reactive oxygen species (ROS) caused by QDs, one study conducted RNAseq transcriptomics to identify a possible mechanism of toxicity. A notable downregulation of cadherin, integrin, and Rho GTPase was discerned. This study concluded that CdSe/ZnS QDs may induce toxicity by dysregulating cytoskeletal component regulation as well as interfering with the adhesion proteins linking cytoskeletal components to the extracellular matrix. This finding prompted us to investigate the expected morphology changes and alteration in cell motility in response to QDs in an attempt to validate the transcriptomic findings.
Motile cells rely on the rapid polymerization and depolymerization of actin filaments to hold and re-shape the cells. Research establishing CdSe/ZnS QDs’ capacity to disrupt actin filament polymerization and depolymerization has been conducted in vitro [28]. Therefore, we hypothesized that QDs will affect the distribution patterns of the actin cytoskeleton, which may negatively impact cell motility. Of the estimated 200 different types of cells in the human body, over 100 different types of cancerous cells are known. As there has been extensive research conducted on quantum dots interacting with HeLa cells, with nearly 1000 papers published in Pubmed, we aimed to compare them to the much less familiar THLE-2 cell line. Currently only one paper exists elucidating the impacts of QDs on THLE-2 [27], but the RNA sequencing data needs more significant validation for cohesive comparison. Thus, we selected THLE-2 and HeLa cells to compare the different cell lines’ subsequent decreases in motile proficiency and connect this to the dysregulation and disruption of actin filament polymerization.

2. Materials and Methods

2.1. Characterization of Cd/Se/ZnS QDs

For this study, we used acid-coated, water-soluble Cd/Se/ZnS QDs (QD620-WS-YY) purchased from NanoOptical Materials Inc. (Carson, CA, USA). We characterized the QDs using emission spectrophotometry and transmission electron microscopy (TEM).
We prepared a QD sample for emission spectra by diluting a stock concentration of 5 mg/mL to 4.3 nM with deionized water. A total of 180 µL was then pipetted into a quartz cuvette. Our emission spectra were measured using a Photon Technology International spectrofluorometer (Birmingham, NJ, USA). We ran emission spectra using a 360 nm excitation wavelength.
We dipped a sterile needle into our QD stock concentration (5 mg/mL) and then dispersed the QD-dipped needle into 1 mL of hexane. A copper grid was then dipped into the hexane solution and allowed to dry overnight. The QD samples were shipped to Arkansas Nano & Bio Materials Characterization Facility for TEM imaging (Fayetteville, AR, USA). An analysis of the TEM images was conducted using ImageJ software (v1.54).

2.2. Quantum Dot Preparation

Our QD stock concentration was 4.3 µM. For all experiments, except XTT, we used bronchial epithelial growth medium (BEGM) to perform a serial dilution to our intended treated QD concentrations. For XTT, we tested QD concentrations that aligned with another paper that tested QDs on THLE-2 cells [27]. Prior to all QD treatments, QDs suspended in BEGM media were sonicated for 30 min to prevent agglomeration.

2.3. Quantum Dot Colocalization with Lysosomes

We used the Biotium LysoView 488 fluorescent label (70067) (Fremont, CA, USA) to visualize the colocalization of our QDs with the lysosomes of THLE-2 and HeLa cells. Following cell quantification, cells were seeded into a glass-bottom 18-well plate. Both THLE-2 and HeLa were seeded at 7000 cells/well and incubated for 24 h. Cells were washed 2× with 1× PBS and then treated with 4.3 nM of QD and incubated for another 24 h. A total of 100 µL of 1× LysoView 488 labeling dye was then added. Following this, cells were incubated for 30 min. We then visualized the fluorescent lysosome structures and quantum dots on a confocal microscope using 488 nm and 561 nm for excitation light for LysoView 488 and QDs, respectively [29,30]. We also used the RFP.SDC and GFP.SDC emission filters on 60× oil (600× overall). We used the JACoP plugin on ImageJ software to calculate Pearson’s correlation coefficient to quantify the colocalization between our QDs and the lysosomes of each cell line.

2.4. Cell Culture

The cell lines used for this study were the THLE-2 and HeLa cell lines. Significant research efforts have been made to understand the effects of QDs on cancerous liver cell lines like HepG2 [31,32,33,34,35], but few studies have expounded the effects of QDs in non-cancerous liver cells [27]. For this reason, we selected non-cancerous cell line THLE-2 to compare it with cancerous cervical cancer line HeLa. THLE-2 cells were cultured in bronchial epithelial growth medium (BEGM) containing the Lonza Walkersville BEGM Bullet Kit (CC-3170, MD) (Lonza, Basel, Switzerland) including bronchial epithelial cell basal medium (BEBM) and the following growth supplements: BPE, 2 mL; Hydrocortisone, 0.5 mL; hEGF, 0.5 mL; Epinephrine, 0.5 mL; Transferrin, 0.5 mL; Insulin, 0.5 mL; Retinoic Acid, 0.5 mL; Triiodothyronine, 0.5 mL; and GA-1000, 0.5 mL. HeLa cells were cultured in Dulbecco’s Modified Eagle Medium (DMEM) (10-013-CM) (Manassas, VA, USA). In addition, 10% fetal bovine serum (FBS) and 1% penicillin and streptomycin were included in both media. Both cell lines were grown in 25 cm2 flasks containing 6 mL of their appropriate media until accordant confluency for each protocol was achieved. Cells were grown in a 37 °C incubator with 5% CO2. To detach cells from their flasks, we removed media, washed them with 1x Phosphate-buffered saline (PBS), and added 1.5 mL of trypsin with EDTA. Following trypsinization, 6 mL of media was added for neutralization. Next, the cells were centrifuged at 400 rpm for 10 min for pellet formation. After removing the supernatant, 6 mL of media was added back in for cell quantification.

2.5. XTT Viability Assay

We used the Biotium XTT Cell Viability Assay Kit (30007) (Fremont, CA, USA) to evaluate the viability of QD-treated THLE-2 and HeLa cells. Following cell quantification, cells were seeded into a 96-well plate. Both THLE-2 and HeLa were seeded at 7000 cells/well and incubated for 24 h. A total of 100 µL of the following concentrations of QDs was treated: 150 nM, 100 nM, 50 nM, 10 nM, and 5 nM. For this experiment, our treated QD concentrations deviated from a 430 nM stock to allow for the direct comparison of XTT assays conducted in a different study [27]. We included a non-treated control as well as 0.1 mM cisplatin and 20% DMSO as positive controls. Each treatment was conducted in quintuplicate. Following another 24 h incubation, the XTT viability kit’s manufacturer’s protocol was followed to create an XTT reagent using a combination of tetrazolium salt XTT and an activating solution in a 200:1 ratio. Each well received 25 µL of XTT reagent and 100 µL of media. The cells were then incubated for 6 h. The XTT reagent was reduced to an orange color, corresponding to cell viability. The colorimetric assay produced absorbance values read by a BioTek ELx880 plate reader (Agilent, Santa Clara, CA, USA) set to the delta wavelength range (450–630 nm). Values were then analyzed using GraphPad Prism9 software (CA).

2.6. Confocal Visualization of THLE-2 Attachment Protrusions

We used a confocal microscope to evaluate morphological changes induced in THLE-2 following QD treatment. Following cell quantification, cells were seeded into a glass-bottom 18-well plate. THLE-2 cells were seeded at 7000 cells/well and incubated for 24 h. Using a serial dilution, 100 µL of the following concentrations of QDs was treated: 430 nM, 43 nM, 4.3 nM, 0.43 nM, and 0.043 nM. We also included a non-treated control (NTC). Each treatment was conducted in triplicate. Following 24 h incubation, cells were washed 2× with 1× PBS. Cell morphology was then analyzed on the DIC channel of an Olympus IX81 microscope with a 10x objective lens (100× overall) and a 60× oil lens (600× overall). We analyzed the average length of the treated and NTC cells. Quantitative analysis was used to further analyze changes in cell morphology. Briefly, all cells maintaining attachment protrusions were considered to be morphologically normal, whereas cells without attachment protrusions were considered to be morphologically abnormal. The percent of each treatment’s change in cell morphology was then analyzed using Dunnett’s multiple comparison test in GraphPad Prism9 software (CA).

2.7. Actin Filament Fluorescence Microscopy

We used the Thermo Fisher Alexa Fluor 488 Phalloidin labeling probe (A12379) (Waltham, MA, USA) to visualize the changes in filamentous actin in THLE-2 and HeLa cells following QD treatment. Following cell quantification, cells were seeded into a glass-bottom 18-well plate. Both THLE-2 and HeLa were seeded at 7000 cells/well and incubated for 24 h. Cells were washed 2× with 1× PBS and then treated with 4.3 nM of QDs and incubated for another 24 h. We followed the Invitrogen phalloidin user manual to fix, permeabilize, and stain actin filaments. Cells were washed 2× with 1× PBS. To fix cells, 23 µL of 16% paraformaldehyde and 77 µL 1× PBS was added to each well and allowed to sit for 15 min at room temperature. Cells were then washed 2× with 1× PBS. To permeabilize cells, 0.1% Triton-X-100 was added to each well and allowed to sit for 15 min at room temperature. Cells were then washed 2× with 1× PBS. A total of 4.5 µL of the 400× stock phalloidin solution was added to a 15 µL falcon tube containing 792 µL 1× PBS and 8 µL of 3.4 mg/mL bovine serum albumin. Cells were then incubated for 60 min and then washed 2× with 1× PBS. We visualized the fluorescent actin filaments on a confocal microscope using 488 nm and 561 nm for excitation and emission lasers, respectively. We also used the RFP.SDC and GFP.SDC channels on 60× oil (600× overall). We analyzed an optical section of the 3D capture images taken to confirm the internalization of QDs. Quantitative analysis was used to analyze distribution changes in F-actin. Cells with extensive networks of linear actin filaments were classified as unchanged. Cells with diminished and disfigured actin networks were classified as abnormal. We quantified the change in F-actin arrangement using Tukey’s multiple comparison test in GraphPad Prism9 software.

2.8. Talin Adhesion Protein Fluorescence Microscopy

We used the CellLight Talin-GFP fluorescent protein labeling reagent (C10611) (Waltham, MA, USA) to visualize the changes in talin, a focal adhesion protein. Following cell quantification, cells were seeded into a glass-bottom 18-well plate. THLE-2 cells were seeded at 7000 cells/well and incubated for 24 h. Cells were then washed 2× with 1× PBS. A total of 14 µL of Celllight talin-GFP was added to each well and incubated for 24 h. Cells were washed 2× with 1× PBS and then treated with 4.3 nM of QDs and incubated for another 24 h. Before visualization, cells were washed 2× with 1× PBS. We used a confocal fluorescent microscope to image cells using the 488 nm and 561 nm excitation/emission wavelength laser set. We counted the number of fluorescent talin-GFP-carrying structures present in a 100 µm2 perimeter. We quantified the number of focal adhesions using Tukey’s multiple comparison test in GraphPad Prism9 software.

2.9. Migration Assay

We used confocal microscopy to evaluate changes in migration ability in QD-treated THLE-2 and HeLa cells. Following cell quantification, cells were seeded into an ibidi Culture Insert 2 Well µ-dish (81176) (Fitchburg, WI, USA). Both THLE-2 and HeLa were seeded at 7000 cells/well and incubated for 24 h with 1 mL of media surrounding the well insert. Cells were washed 2× with 1× PBS and then treated with 4.3 nM of QDs and incubated for another 24 h. Cells were washed 2× with 1× PBS and then resubmerged in media. A total of 1 mL of media was also added to the area surrounding the well insert. We then slowly removed the well insert using sterile forceps. We imaged both cell lines’ gap length every 3 h over the span of 24 h. We quantified the change in gap length using Šidák’s multiple comparison test in GraphPad Prism9 software.

2.10. Statistical Analysis

Statistical analysis was conducted using GraphPad Prism 9 for all experiments. When evaluating XTT and morphology, we ran a One-way ANOVA test to compare the treated columns to the control. We ran a Two-way ANOVA when evaluating abnormal actin and migration gap distance between groups. We also used an unpaired t test to evaluate the frequency between the control and QD-treated groups’ talin-GFP-carrying focal adhesion.

3. Results and Discussion

3.1. Characterization of Quantum Dots

In this study we used Cd/Se/ZnS QDs containing a surface thiol conjugated to carboxylic acid and solubilized in water. The emission peak of our QD sample read 620 nm (Figure 1a). We also analyzed our QD TEM images to find that the average size of our QDs was 7.259 ± 1.072 nm (Figure 1b,c). This was 1.059 nm larger than the 6.2 nm size stated by the manufacturer.

3.2. Colocalization of QDs and Lysosomes in THLE-2 and HeLa Cells

To confirm the internalization of our QDs, we conducted a colocalization experiment to determine the location of our QDs within the cell. In THLE-2 cells, colocalization between QDs and lysosomes was evidenced by a Pearson’s correlation coefficient of 0.429 ± 0.0693. HeLa cells had a higher degree of colocalization as the respective Pearson’s correlation coefficient was 0.761 ± 0.0871. Figure 2 confirms the internalization of QDs into both cell lines. As the colocalization between lysosomes and QDs, indicated by Pearson’s correlation coefficient, is higher in HeLa cells than THLE-2 cells, we believe this may suggest that HeLa cells internalize QDs at a faster rate than THLE-2 cells. This could lead to a greater accumulation of QDs within the lysosome. We also hypothesize that HeLa cells may remove QDs via the lysosome pathway at a higher rate than THLE-2 cells. Our hypothesis is further bolstered by a similar study that indicated that QDs are commonly removed from cells via lysosomal exocytosis [36].

3.3. Impacts of Different QDs on THLE-2 and HeLa Cell Viability

We used an XTT viability assay to evaluate the cytotoxic effects that differing concentrations of Cd/Se/ZnS QDs have on THLE-2 cells and HeLa cells. Our results indicate that QDs decrease THLE-2 cell viability in a dose-dependent manner (Figure 3a). Our lowest treated concentration of QD 5 nM showed the smallest decrease in THLE-2 viability though it still indicated statistical significance. As all concentrations of treated QDs induced significant decreases in THLE-2 cell viability, an IC50 value could not be calculated on SlideBook Prism9. In the following fluorescent microscopy experiments, we treated cells with 4.3 nM QD as 5 nM exhibited the smallest decrease in cell viability. QDs produced contrasting results in HeLa cells as no changes in viability were seen at any concentrations (Figure 3b).
A prior study looked at THLE-2 cell viability compared to cancerous liver cell HepG2 viability following CdSe/ZnS QD exposure [27]. The authors found that QD concentrations as low as 50 nM did not cause a decrease in THLE-2 cell viability. Our THLE-2 cells lost significant viability at one tenth of this QD concentration. The stark difference between their results and ours in the current report might be due to the different size of QDs (their QDs are 6 nm in size and emit 450 nm, and the QDs used in this study are 8 nm and emit 620 nm) and different cell lines used between the two reports. Interestingly, our XTT results indicate that THLE-2 cells are significantly more sensitive to Cd/Se/ZnS QDs than HeLa cells as we saw no decrease in HeLa cell viability at any QD concentration. Somewhat consistent with this finding, a recent report demonstrated that HeLa cells are more resistant to red CdSe/ZnS QDs than ML-1 thyroid cancer cells. The authors proposed that the observed resistance in HeLa cells can be attributed to the faster metabolic rate or cell proliferation rate in HeLa. Considering that THLE-2’s doubling time is substantially slower (at least 4–5 times) than that of HeLa cells, our interpretation of the cell viability defects observed in THLE-2 cells is related to their slow cell proliferation rate [25]. Consistently, some researchers have found that cervical cancer in mice incurs no decreases in cell viability even at high QD concentrations [37]. Nevertheless, other studies have found that QD-treated HeLa cells cause a decrease in viability,; however, some of the QD concentrations that induced significant decreases in viability (168 nM and 196 nM) were higher than our highest treated QD concentration [24,25]. If we increased the range of QD concentrations treated, it is possible that we might have seen a similar trend in HeLa cell viability.
As our results strongly indicate that QDs are toxic to in vitro liver epithelial cells, we believe that this may provide insight into the possible effects of QDs on in vivo experiments. As the small size of QDs allows for easy cellular uptake, investigating the big picture effects of bioaccumulation within an organism’s liver is of high interest. Similar studies confirm our hypothesis that QDs entering an organism’s circulatory system result in the bioaccumulation of QDs in the liver as the liver’s primary function is to detoxify blood [38,39,40,41]. The uptake of QDs in our THLE-2 cells resulted in cytotoxicity, signifying dysfunctional intracellular processes. Evidence of disturbances within the cell is commonly known to manifest in increased reactive oxygen species (ROS) levels [42]. We hypothesize that if QDs induce ROS, an inflammatory response may occur locally in the liver, resulting in changes in metabolism as well as permanent hepatic tissue damage. Long-term exposure to QDs may result in a perpetuated inflammatory response that can lead to detrimental chronic liver diseases like non-alcoholic fatty liver disease or hepatitis [43].

3.4. Confocal Visualization Revealed Morphological Changes in THLE-2 Cells

We found that QDs cause significant changes in THLE-2 length and cell morphology. With the average length of an NTC THLE-2 cell being 120.34 µm (Figure 4a), exposure to our highest QD concentrations (430 nM) decreased the average cell length to 40.27 µm (Figure 4b). The significant decrease in length was due to the loss of attachment structures projecting from the cell body.
QDs caused THLE-2 cells to become round, characterized by the loss of their spindle-shaped morphology seen in the non-treated cells (Figure 5a). At our smallest QD concentration, 0.043 nM, 46.04 percent of THLE-2 cells lost attachment protrusions (Figure 5b). At our largest QD concentration of 430 nM, 100 percent of THLE-2 cells exhibited a round morphology.
A 1985 study identified three categories of epithelial cell morphology cultured on a 2D substrate: poorly spread, unpolarized but well spread, and polarized and well spread [44]. Poorly spread cells were small and round and had little contact with the substrate. Well-spread but unpolarized cells had a large area contacting the media and multiple weak protrusions around the cell perimeter. Well-spread polarized cells had a large area contacting the media as well as 1–2 strong protrusions, giving the cell an elongated spindle shape. Later studies further expanded upon the morphological changes a cell must undergo before any movement may occur [45,46,47]. When a cell first contacts its substrate, it will have a round morphology until it attaches. It will then flatten as it expands its basal surface. As it continues to spread, protrusion bodies will form as it tests the local environment before finally becoming polarized and moving in a specific direction. We condensed our analysis into two morphology classifications: round with no attachment protrusions and flat with attachment protrusions. We found that QD treatment concentration was positively correlated to the loss of attachment protrusions. Another in vitro study found that CdSe QDs with no functional coating induced similar morphological changes following QD treatment. At 20 nM of QDs, cells lost protrusions from their peripheral membrane [48]. Although they reported no morphological changes induced in concentrations ≤ 10 nM, the common toxic features derived from cadmium-based QDs are similar in that cells lose their distinctive spindle-shaped morphology.

3.5. Fluorescence Microscopy Displayed Redistribution of Actin Filaments

Following our observation of QD-induced morphological changes evidenced by a decrease in the protrusion bodies of a cell, we analyzed the redistribution of actin filaments in THLE-2 and HeLa cells following QD exposure. Three-dimensional microscopy optical sectioning confirmed that QDs were internalized by both cell lines, as well as partially colocalizing to F-actin (Figure 6a,b). Non-treated THLE-2 cells exhibited protrusion bodies containing long fiber-like or stress fiber-like actin filaments (Figure 6c). When treated with 4.3 nM of QDs, THLE-2 cells lost their prominent long stress fibers and gained many short and thin actin filament bundles congregating at the periphery of the cell membrane (Figure 6e). Upon 4.3 nM QD exposure, 68.10% of THLE-2 cells exhibited this abnormal actin redistribution (Figure 6g). NTC HeLa cells also exhibited a similar F-actin distribution, as shown in the NTC THLE-2 cells in Figure 6d. When treated with 4.3 nM of QDs, HeLa cells exhibited less prominent stress fibers as compared with NTC HeLa cells. QD-treated HeLa cells also gained many more short actin bundles near the cell periphery (Figure 6f). Upon 4.3 nM QD exposure, 44.38% of HeLa cells exhibited this change in actin redistribution (Figure 6g).
Upon QD treatment, a significant percentage of THLE-2 and HeLa cells underwent a redistribution of F-actin by losing a substantial amount of stress fibers. Another study also identified similar alterations upon carbon QD exposure [49]. They reported that exposure to 2.5 × 1015 particles/mL of carbon QDs resulted in the partial destruction of the stress fibers, indicated by a 31.35% decrease in F-actin phalloidin intensity, within the breast cancer cell line MCF-7 [49]. Similarly, a study treating human lung cells with 50 to 200 μg/mL silicon QDs observed F-actin redistribution [50]. They found that actin stress fibers were poorly formed and identified the presence of microspikes and membrane ruffling following 24 h of QD incubation. Their imaging also revealed bead-like actin structures not seen in our experiment. This actin structure may be unique to their study due to their high concentration of treated QDs. In light of our findings that many microspike-like structures are peripherally associated with QD-treated cells and that an abundance of microspikes are functionally associated with the loss of directionality of motile cells, we propose that QDs at 4.3 nM are sufficient enough to disrupt the normal behavior of motile cells. Nevertheless, we could not exclude the possibility that the observed spike-like structure was filopodia, which is of interest to be further explored.

3.6. Fluorescence Microscopy Showed Decreased Talin Structures

Talin is a key component in the focal adhesion structures that link the extracellular matrix to the intracellular actin filaments, which allows for the direct mediation of cell adhesion [51]. Talin is also involved in the signal transduction of Rho GTPases [52,53]. In particular, the activation of Rac1 (a specific Rho GTPase) results in changes to a cell’s adhesion and migration abilities [54,55,56]. The earlier sections of this report demonstrated the severe disruption of the actin cytoskeleton, and therefore, we wanted to examine the possibility that the integrity of focal adhesion is crippled in response to QDs. For this we analyzed the abundance of the focal adhesion marked with talin-GFP in non-treated and QD-treated THLE-2 cells. Following QD treatment, we saw a decrease in the average number of talin-carrying focal adhesion in the ROI (a 100 µm2 box in the image) (Figure 7). NTC cells had an average of 24.61 focal adhesions in the defined perimeter. THLE-2 cells treated with 4.3 nM of QDs had an average of 16.30 focal adhesions in the defined perimeter. We propose that the reduced number of focal adhesions is linked to the dysregulation and disruption of the actin cytoskeleton in QD-treated cells. Although QDs can directly interact with talin to disturb the structural integrity of focal adhesions, at this time, there is no supporting evidence for this notion from our study nor in the literature. Corresponding studies from Harris et al. showed that 61 nM of Cd/Se/ZnS QDs (different size) caused a downregulation of genes regulating Rho GTPase and integrin in THLE-2 cells, reflecting that the reduced number of focal adhesions might be due to the low expression of talin [27].

3.7. QDs Decreased Cell Migration in THLE-2 and HeLa

We used a 2-well migration assay to assess the ability of THLE-2 and HeLa cells to migrate following QD exposure. Both cell lines incurred a substantial decrease in motility upon QD exposure. A total of 3 h after barrier removal, NTC THLE-2 cells moved an average of 55.11 µm (Figure 8b). This was significantly more than THLE-2 cells treated with 4.3 nM of QDs, which moved an average of 15.34 µm 3 h post barrier removal. The difference in gap closure continued to increase, and at 24 h post barrier removal, NTC THLE-2 cells completely migrated into the gap, whereas QD-treated THLE-2 cells maintained an average gap length of 161.99 µm. The average velocity of non-treated THLE-2 cells was 27.85 µm/h, whereas THLE-2 cells exposed to QDs had an average velocity of 17.24 µm/h (Figure 8c).
HeLa cells also underwent a significant change in gap closure compared to the NTC at 3 h. A total of 3 h after barrier removal, NTC HeLa cells moved an average of 48.56 µm (Figure 9b). This was significantly more than QD-treated HeLa cells. After 3 h, HeLa cells treated with 4.3 nM of QDs only moved an average of 26.6 µm. The difference in gap closure appeared to peak at 18 h post barrier removal as only non-treated HeLa cells completely closed the gap. At 24 h both non-treated and QD-treated HeLa cells closed the gap completely. The average velocity of non-treated HeLa cells was 41.47 µm/h, whereas HeLa cells exposed to QDs had an average velocity of 30.01 µm/h (Figure 8c).
We noticed that QD-treated HeLa, non-treated HeLa, and non-treated THLE-2 cells displayed a peak in migration speed at 18 h post barrier removal, followed by a subsequent decrease in migration speed (Figure 8c and Figure 9c). We attribute this decrease in migration speed to a decrease in the available area in which cells could migrate. This would be especially prominent if QD-treated HeLa, non-treated HeLa, and non-treated THLE-2 cells fully closed the gap prior to the 24 h mark. QD-treated THLE-2 cells further prove this hypothesis as these cells never fully closed their gap and thus never ran out of available area to migrate within. This was moreover indicated by no dip being seen in migration speed during the 18 to 24 h time period.
QD-treated THLE-2 and HeLa cells had a decreased ability to migrate when compared to their non-treated controls. Similarly, a recent study treated breast cancer cells (MCF-7 and MDA-MB-231) with carbon quantum dots (CQDs) and found that CQDs gave rise to similar impeded cell migration [57]. At 48 h, non-treated MFC-7 cells closed 94% of their gap, while CQD-treated MFC-7 cells only closed 13% of their gap. Another study established analogous results when they treated U87MG glioma cells with 100 nM of gold quantum dots (GQDs), significantly reducing the migration and invasion abilities of the U87MG glioma cells at 24 h [58]. The concentration of GQDs needed to significantly reduce the migration ability of U87MG glioma cells was much higher than our 4.3 nM concentration of Cd/Se/ZnS QDs. We suggest that the vulnerability of our cells may be due in part to cadmium ions leaking from the core and past the ZnS shell. When rats were exposed to Cd/Se QDs, free Cd ions were selectively identified in their liver and kidneys, demonstrating the prominent leakage of Cd/Se QDs [59]. Some studies find that a ZnS shell prevents leakage, thereby mitigating the toxicity of the Cd QD core [20,21,60,61,62]; however, others find that the integrity of QD shell coatings decreases once in a biological system [63]. For example, the endocytosis of Cd/Se/ZnS QDs still released Cd ions as the low pH of endosomes degraded the shells [64].
Both QD-treated THLE-2 and HeLa cells had a decrease in average cell velocity as both were roughly 10 µm/h slower than that of their non-treated controls. A recent study found that CQDs have also been shown to decrease breast cancer cell velocity [57]. Based on their graph, non-treated MFC-7 cells had an average velocity of 24 µm/h, and CQD-treated MFC-7 cells had an average velocity of 16.5 µm/h, indicating that QDs decreased the average cell velocity by 7.5 µm/h. This decrease in average velocity is 25 percent less than our average velocity of 10 µm/h, which we attribute to the absence of toxic heavy metal ions leaking from the CQDs [65,66,67].
When comparing non-treated THLE-2 cells to HeLa cells, we observed that HeLa cells migrate more efficiently as they fully closed the gap in 18 h, whereas THLE-2 cells took 24 h to close the gap. QD-treated cells in both cell lines follow this trend. QD-treated THLE-2 never closed the gap, while HeLa cells treated with QDs closed the gap after 24 h. When comparing the non-treated and QD-treated average velocities between the two cell lines, HeLa cells have a higher average velocity in both scenarios. This data suggests that HeLa cells possess an inherent increased ability to migrate faster than THLE-2. HeLa cells have a duplication time of 18–24 h as well as increased migration abilities due to their metastatic state [68]. In contrast, THLE-2 cells have a generation time 2× to 4× that of HeLa (2–4 days) [69]. We suggest that HeLa cells’ fast doubling time was able to partially compensate for the QD-treated cells’ decreased ability to migrate, though ultimately QD-treated HeLa cells still closed the barrier gap slower than non-treated HeLa cells. A transcriptomic study carried out by Harris et al. found that Cd/Se/ZnS QDs downregulate the Rho GTPase pathway in THLE-2 cells [27]. It is likely that this impedes the downstream signal cascade, resulting in the disruption of actin polymerization in cellular protrusions. We propose that the QD downregulation of this pathway may explain the pronounced decrease in THLE-2 cells’ migration ability. In the future, an evaluation of HeLa cells’ transcriptional changes upon QD exposure would confirm whether this pathway was downregulated to a similar degree.

4. Conclusions and Future Directions

As advancements are made in the biomedical field’s application of QDs, so should our efforts to understand the consequential intracellular effects of these nanomaterials. In this study, we assessed the toxicity of Cd/Se/ZnS QDs, as well as the collateral morphological consequences. Our experiments reveal that Cd/Se/ZnS more drastically reduced cell viability, actin stress fiber development, and migration ability in THLE-2 cells than HeLa cells. When explaining our findings, we base much of our rationale off of THLE-2 transcriptomic research [27]. In the future, an assessment of the regulatory changes in HeLa cells will permit a more thorough comparison between cell lines. We also believe that our findings strongly indicate a need for a safer alternative. One promising alternative is carbon quantum dots. Future studies should focus on assessing whether the alternatives to heavy metal QDs have lesser effects on cell viability while still possessing similar chemical and drug conjugation potential.

Author Contributions

Conceptualization, M.M., A.C. and K.K.; methodology, M.M. and A.C.; software, M.M.; validation, M.M., A.C. and K.K.; formal analysis, M.M.; investigation, M.M. and A.C.; resources, K.K.; data curation, M.M. and A.C.; writing—original draft preparation, M.M.; writing—review and editing, K.K.; visualization, M.M., A.C. and K.K.; supervision, K.K.; project administration, K.K.; funding acquisition, K.K. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by Roy Blunt Professorship funds at Missouri State University.

Data Availability Statement

All data are contained within this article.

Acknowledgments

We would like to show appreciation to K.K. for his support and guidance during every step of this project. We would also like to thank Emma Braun for her assistance in QD characterization. Lastly, we want to thank Missouri State University for providing the space and equipment necessary for this research project.

Conflicts of Interest

The authors declare no conflicts of interest.

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Figure 1. Emission spectra of QDs exposed to 360 nm excitation wavelength (a). Frequency distribution of QD size (b) and TEM image of Cd/Se/ZnS QDs (c). Scale bar in (c) is 2 nm.
Figure 1. Emission spectra of QDs exposed to 360 nm excitation wavelength (a). Frequency distribution of QD size (b) and TEM image of Cd/Se/ZnS QDs (c). Scale bar in (c) is 2 nm.
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Figure 2. Confocal visualization of QDs in THLE-2 cell on RFP.SDC channel (a) as well as lysosomes on GFP.SDC channel (b) demonstrating colocalization when merged (c). Confocal visualization of QDs in HeLa cell on RFP.SDC channel (d) as well as lysosomes on GFP.SEC channel (e) demonstrating colocalization when merged (f). Scale bars in (a–f) are 5 µm.
Figure 2. Confocal visualization of QDs in THLE-2 cell on RFP.SDC channel (a) as well as lysosomes on GFP.SDC channel (b) demonstrating colocalization when merged (c). Confocal visualization of QDs in HeLa cell on RFP.SDC channel (d) as well as lysosomes on GFP.SEC channel (e) demonstrating colocalization when merged (f). Scale bars in (a–f) are 5 µm.
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Figure 3. XTT viability assay of THLE-2 cells exposed to 150 nM, 100 nM, 50 nM, 10 nM, and 5 nM of QDs (a) and QDs exposed HeLa cells (b). Statistical analysis was conducted using Dunnett’s multiple comparison test, * p < 0.05 and **** p < 0.0001.
Figure 3. XTT viability assay of THLE-2 cells exposed to 150 nM, 100 nM, 50 nM, 10 nM, and 5 nM of QDs (a) and QDs exposed HeLa cells (b). Statistical analysis was conducted using Dunnett’s multiple comparison test, * p < 0.05 and **** p < 0.0001.
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Figure 4. Frequency distribution of NTC THLE-2 cell length (a) and frequency distribution of 430 nM QD-treated THLE-2 cell length (b).
Figure 4. Frequency distribution of NTC THLE-2 cell length (a) and frequency distribution of 430 nM QD-treated THLE-2 cell length (b).
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Figure 5. DIC.SDC channel confocal images of THLE-2 cells exposed to 430 nM, 43 nM, 4.3 nM, 0.43 nM, and 0.043 nM of QDs (a). Percent of THLE-2 cells become round following QD treatment (b). Statistical analysis was conducted using Dunnett’s multiple comparison test, * p < 0.05, ** p < 0.01, and **** p < 0.0001. Scale bars in (a) are 50 µm.
Figure 5. DIC.SDC channel confocal images of THLE-2 cells exposed to 430 nM, 43 nM, 4.3 nM, 0.43 nM, and 0.043 nM of QDs (a). Percent of THLE-2 cells become round following QD treatment (b). Statistical analysis was conducted using Dunnett’s multiple comparison test, * p < 0.05, ** p < 0.01, and **** p < 0.0001. Scale bars in (a) are 50 µm.
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Figure 6. Three-dimensional confocal visualization using GFP.SDC and RFP.SDC channels, confirming internalization of QDs. Pink box: Full zoom of QDs colocalizing with actin filaments in THLE-2 cell (a) Blue box: Full zoom of QDs colocalizing with actin filaments in HeLa cell (b). Actin filament distribution in NTC THLE-2 cell (c) and NTC HeLa cell (d). Actin filament distribution in 4.3 nM QD-treated THLE-2 cell (e) and 4.3 nM QD-treated HeLa cell (f). Percent of treated and non-treated THLE-2 and HeLa cells lacking well-developed actin stress fibers. (g). Statistical analysis was conducted using Tukey’s multiple comparison test * p < 0.05 **** p < 0.0001. Lower left scale bars in (a–f) are 20 µm. Scale bars in pink and blue zoom boxes are 2 µm.
Figure 6. Three-dimensional confocal visualization using GFP.SDC and RFP.SDC channels, confirming internalization of QDs. Pink box: Full zoom of QDs colocalizing with actin filaments in THLE-2 cell (a) Blue box: Full zoom of QDs colocalizing with actin filaments in HeLa cell (b). Actin filament distribution in NTC THLE-2 cell (c) and NTC HeLa cell (d). Actin filament distribution in 4.3 nM QD-treated THLE-2 cell (e) and 4.3 nM QD-treated HeLa cell (f). Percent of treated and non-treated THLE-2 and HeLa cells lacking well-developed actin stress fibers. (g). Statistical analysis was conducted using Tukey’s multiple comparison test * p < 0.05 **** p < 0.0001. Lower left scale bars in (a–f) are 20 µm. Scale bars in pink and blue zoom boxes are 2 µm.
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Figure 7. Fluorescence microscopy of talin-carrying focal adhesion structures in THLE-2 cells using GFP and TxRed channels with 488 and 561 nm excitation wavelengths (a). Number of focal adhesions in NTC and 4.3 nM QD-treated THLE-2 cells (b). Statistical analysis of average number of focal adhesions was conducted using unpaired t test. *** p < 0.005. Scale bars in (a) are 20 µm.
Figure 7. Fluorescence microscopy of talin-carrying focal adhesion structures in THLE-2 cells using GFP and TxRed channels with 488 and 561 nm excitation wavelengths (a). Number of focal adhesions in NTC and 4.3 nM QD-treated THLE-2 cells (b). Statistical analysis of average number of focal adhesions was conducted using unpaired t test. *** p < 0.005. Scale bars in (a) are 20 µm.
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Figure 8. DIC.SDS channel confocal images of gap 0, 3, 6, 9, 18, and 24 h post barrier removal in THLE-2 NTC and QD-treated cells (a). Average gap length in THLE-2 NTC and QD-treated cells at 0, 3, 6, 9, 18, and 24 h post barrier removal (b). Average velocity of THLE-2 NTC and QD-treated cells at 0, 3, 6, 9, 18, and 24 h post barrier removal (c). Gap length analyzed using Šidák’s multiple comparison test. ** p < 0.01 and **** p < 0.0001. Scale bars in (a) are 50 µm.
Figure 8. DIC.SDS channel confocal images of gap 0, 3, 6, 9, 18, and 24 h post barrier removal in THLE-2 NTC and QD-treated cells (a). Average gap length in THLE-2 NTC and QD-treated cells at 0, 3, 6, 9, 18, and 24 h post barrier removal (b). Average velocity of THLE-2 NTC and QD-treated cells at 0, 3, 6, 9, 18, and 24 h post barrier removal (c). Gap length analyzed using Šidák’s multiple comparison test. ** p < 0.01 and **** p < 0.0001. Scale bars in (a) are 50 µm.
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Figure 9. Confocal images on DIC.SDS channel of gap 0, 3, 6, 9, 18, and 24 h post barrier removal in NTC and QD-treated cells (a). Average gap length in HeLa NTC and QD-treated cells at 0, 3, 6, 9, 18, and 24 h post barrier removal (b). Average velocity of HeLa NTC and QD-treated cells at 0, 3, 6, 9, 18, and 24 h post barrier removal (c). Gap length analyzed using Šidák’s multiple comparison test. *** p < 0.001 and **** p < 0.0001. Scale bars in (a) are 50 µm.
Figure 9. Confocal images on DIC.SDS channel of gap 0, 3, 6, 9, 18, and 24 h post barrier removal in NTC and QD-treated cells (a). Average gap length in HeLa NTC and QD-treated cells at 0, 3, 6, 9, 18, and 24 h post barrier removal (b). Average velocity of HeLa NTC and QD-treated cells at 0, 3, 6, 9, 18, and 24 h post barrier removal (c). Gap length analyzed using Šidák’s multiple comparison test. *** p < 0.001 and **** p < 0.0001. Scale bars in (a) are 50 µm.
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Metcalf, M.; Chand, A.; Kim, K. Quantum Dots Affect Actin Cytoskeleton Reorganization, Resulting in Impaired HeLa and THLE-2 Cell Motility. Micro 2025, 5, 29. https://doi.org/10.3390/micro5020029

AMA Style

Metcalf M, Chand A, Kim K. Quantum Dots Affect Actin Cytoskeleton Reorganization, Resulting in Impaired HeLa and THLE-2 Cell Motility. Micro. 2025; 5(2):29. https://doi.org/10.3390/micro5020029

Chicago/Turabian Style

Metcalf, Mileah, Abhishu Chand, and Kyoungtae Kim. 2025. "Quantum Dots Affect Actin Cytoskeleton Reorganization, Resulting in Impaired HeLa and THLE-2 Cell Motility" Micro 5, no. 2: 29. https://doi.org/10.3390/micro5020029

APA Style

Metcalf, M., Chand, A., & Kim, K. (2025). Quantum Dots Affect Actin Cytoskeleton Reorganization, Resulting in Impaired HeLa and THLE-2 Cell Motility. Micro, 5(2), 29. https://doi.org/10.3390/micro5020029

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