# Dynamic Effective Elasticity of Melanoma Cells under Shear and Elongational Flow Confirms Estimation from Force Spectroscopy

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## Abstract

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## 1. Introduction

^{®}CRL-1619™) cell line is a human melanoma cell line initiated through explant culture of a solid tumor from a 54-year-old female and has been used to, for example, obtain paracrine factors for the prolonged culture of mesenchymal stromal cells [7]. On that basis, we study the elasticity of A375 melanoma cells using force spectroscopy (FS) and microfluidic deformation analysis.

## 2. Methods

#### 2.1. Cell Culture

^{®}CRL-1619™) being studied are human malignant melanoma cells. They were cultivated under standard conditions of 37 °C and 5% CO2 in DMEM (30-2002, ATCC, Manassas, VA, USA) containing 10% FBS (Bio&SELL GmbH, Feucht bei Nürnberg, Germany) and 1% Penicillin-Streptomycin (Sigma–Aldrich Chemie GmbH, Steinheim, Germany).

#### 2.2. Force Spectroscopy

#### 2.3. Microfluidic Method Microchannel Design and Measurement Setup

#### 2.4. Statistics

## 3. Results

#### 3.1. Elasticity of Melanoma Cells—Force Spectroscopy

_{day, no tip}= −0.08), and with the tip we find a low ICC of ICC

_{day, tip}= 0.2. Comparing the scattering of the values within one cell to the differences between cells, however, results in high ICC values (ICC

_{cell, no tip}= 0.84, ICC

_{cell, tip}= 0.66). In conclusion, the values correlate strongly within a cell, and therefore only mean values over each cell are summarized in the box plot. The low ICC

_{day}values justify treating each cell as an independent measurement. This means that the biological variation is the main origin of the broad range of obtained results for the Young´s modulus E. The lower correlation and wider distribution for the results with the tip might arise for two reasons: If the cell is penetrated, higher values may occur because the underlying harder substrate contributes to the effective elasticity [29]. In addition, the inhomogeneity of the cell has a greater influence on the result with a smaller contact area.

#### 3.2. Elasticity of Melanoma Cells—Microfluidic Method

#### 3.3. Limitations of the Method

## 4. Discussion

#### Comparison of Measurement Techniques

_{0}(200 ms) = 101 ± 8 Pa and G

_{0}(5 ms) = 590 ± 5 Pa for different time constants. For validation, they refer to optical stretcher experiments, which lead to values in the range of 70–100 Pa. Applying their microfluidic method to GBM tumor initiating cells, they find G

_{0}= 440 ± 0.03 Pa, and report AFM results of G

_{0}= 800–900 Pa [19]. Chen et al., who also developed a microfluidic method, which is based on measuring the changes in pressure while a cell passes a stenosis, provide another set of different cell lines and methods. The results for K562 leukemia cells are 64 Pa from the microfluidic method, 90 Pa from micropipette aspiration and 400 Pa measured by AFM. The same three measurement methods do not show the same systematics for endothelial cells: 383 Pa from the microfluidic method, 100–400 Pa from micropipette aspiration and 700–3000 Pa using AFM [18].

## 5. Conclusions

## Supplementary Materials

## Author Contributions

## Funding

## Institutional Review Board Statement

## Informed Consent Statement

## Data Availability Statement

## Acknowledgments

## Conflicts of Interest

## References

- Dive, C.; Brady, G. SnapShot: Circulating Tumor Cells. Cell
**2017**, 168, 742–742.e1. [Google Scholar] [CrossRef] - Marsavela, G.; Aya-Bonilla, C.A.; Warkiani, M.E.; Gray, E.S.; Ziman, M. Melanoma circulating tumor cells: Benefits and challenges required for clinical application. Cancer Lett.
**2018**, 424, 1–8. [Google Scholar] [CrossRef] [PubMed] - Kilgour, E.; Rothwell, D.G.; Brady, G.; Dive, C. Liquid Biopsy-Based Biomarkers of Treatment Response and Resistance. Cancer Cell
**2020**, 37, 485–495. [Google Scholar] [CrossRef] - Gorges, K.; Wiltfang, L.; Gorges, T.M.; Sartori, A.; Hildebrandt, L.; Keller, L.; Volkmer, B.; Peine, S.; Babayan, A.; Moll, I.; et al. Intra-Patient Heterogeneity of Circulating Tumor Cells and Circulating Tumor DNA in Blood of Melanoma Patients. Cancers
**2019**, 11, 1685. [Google Scholar] [CrossRef] [Green Version] - Thomas, G.; Burnham, N.A.; Camesano, T.A.; Wen, Q. Measuring the mechanical properties of living cells using atomic force microscopy. J. Vis. Exp.
**2013**, e50497. [Google Scholar] [CrossRef] [PubMed] [Green Version] - Mondadori, C.; Crippa, M.; Moretti, M.; Candrian, C.; Lopa, S.; Arrigoni, C. Advanced Microfluidic Models of Cancer and Immune Cell Extravasation: A Systematic Review of the Literature. Front. Bioeng. Biotechnol.
**2020**, 8, 1–19. [Google Scholar] [CrossRef] [PubMed] - Kucerova, L.; Matuskova, M.; Hlubinova, K.; Altanerova, V.; Altaner, C. Tumor cell behaviour modulation by mesenchymal stromal cells. Mol. Cancer
**2010**, 9, 129. [Google Scholar] [CrossRef] [Green Version] - Wu, P.-H.; Aroush, D.R.-B.; Asnacios, A.; Chen, W.-C.; Dokukin, M.E.; Doss, B.L.; Durand-Smet, P.; Ekpenyong, A.; Guck, J.; Guz, N.V.; et al. A comparison of methods to assess cell mechanical properties. Nat. Methods
**2018**, 15, 491–498. [Google Scholar] [CrossRef] [PubMed] - Hao, Y.; Cheng, S.; Tanaka, Y.; Hosokawa, Y.; Yalikun, Y.; Li, M. Mechanical properties of single cells: Measurement methods and applications. Biotechnol. Adv.
**2020**, 45, 107648. [Google Scholar] [CrossRef] - Wu, P.-H.; Hale, C.M.; Chen, W.-C.; Lee, J.S.H.; Tseng, Y.; Wirtz, D. High-throughput ballistic injection nanorheology to measure cell mechanics. Nat. Protoc.
**2012**, 7, 155–170. [Google Scholar] [CrossRef] [Green Version] - Oh, M.-J.; Kuhr, F.; Byfield, F.; Levitan, I. Micropipette aspiration of substrate-attached cells to estimate cell stiffness. J. Vis. Exp.
**2012**, e3886. [Google Scholar] [CrossRef] [PubMed] [Green Version] - Hochmuth, R.M. Micropipette aspiration of living cells. J. Biomech.
**2000**, 33, 15–22. [Google Scholar] [CrossRef] - Hu, S.; Wang, R.; Tsang, C.M.; Tsao, S.W.; Sun, D.; Lam, R.H.W. Revealing elasticity of largely deformed cells flowing along confining microchannels. RSC Adv.
**2018**, 8, 1030–1038. [Google Scholar] [CrossRef] [Green Version] - Kuznetsova, T.G.; Starodubtseva, M.; Yegorenkov, N.; Chizhik, S.A.; Zhdanov, R.I. Atomic force microscopy probing of cell elasticity. Micron
**2007**, 38, 824–833. [Google Scholar] [CrossRef] [PubMed] - Streppa, L.; Ratti, F.; Goillot, E.; Devin, A.; Schaeffer, L.; Arneodo, A.; Argoul, F. Prestressed cells are prone to cytoskeleton failures under localized shear strain: An experimental demonstration on muscle precursor cells. Sci. Rep.
**2018**, 8, 8602. [Google Scholar] [CrossRef] [Green Version] - Amiri, A.; Hastert, F.D.; Heim, L.-O.; Dietz, C. Reliability of cancer cell elasticity in force microscopy. Appl. Phys. Lett.
**2020**, 116, 083701. [Google Scholar] [CrossRef] - Virumbrales-Muñoz, M.; Paz-Artigas, L.; Ciriza, J.; Alcaine, C.; Espona-Noguera, A.; Doblaré, M.; Del Burgo, L.S.; Ziani, K.; Pedraz, J.L.; Fernández, L.; et al. Force Spectroscopy Imaging and Constriction Assays Reveal the Effects of Graphene Oxide on the Mechanical Properties of Alginate Microcapsules. ACS Biomater. Sci. Eng.
**2021**, 7, 242–253. [Google Scholar] [CrossRef] - Chen, Z.; Zhu, Y.; Xu, D.; Alam, M.; Shui, L.; Chen, H. Cell elasticity measurement using a microfluidic device with real-time pressure feedback. Lab a Chip
**2020**, 20, 2343–2353. [Google Scholar] [CrossRef] - Guillou, L.; Dahl, J.; Lin, J.-M.G.; Barakat, A.I.; Husson, J.; Muller, S.J.; Kumar, S. Measuring Cell Viscoelastic Properties Using a Microfluidic Extensional Flow Device. Biophys. J.
**2016**, 111, 2039–2050. [Google Scholar] [CrossRef] [Green Version] - Dudani, J.S.; Gossett, D.R.; Tse, H.T.K.; Di Carlo, D. Pinched-flow hydrodynamic stretching of single-cells. Lab Chip
**2013**, 13, 3728–3734. [Google Scholar] [CrossRef] - Armistead, F.J.; De Pablo, J.G.; Gadêlha, H.; Peyman, S.A.; Evans, S.D. Cells Under Stress: An Inertial-Shear Microfluidic Determination of Cell Behavior. Biophys. J.
**2019**, 116, 1127–1135. [Google Scholar] [CrossRef] [PubMed] [Green Version] - Mietke, A.; Otto, O.; Girardo, S.; Rosendahl, P.; Taubenberger, A.; Golfier, S.; Ulbricht, E.; Aland, S.; Guck, J.; Fischer-Friedrich, E. Extracting Cell Stiffness from Real-Time Deformability Cytometry: Theory and Experiment. Biophys. J.
**2015**, 109, 2023–2036. [Google Scholar] [CrossRef] [PubMed] [Green Version] - Otto, O.; Rosendahl, P.; Mietke, A.; Golfier, S.; Herold, C.; Klaue, D.; Girardo, S.; Pagliara, S.; Ekpenyong, A.; Jacobi, A.; et al. Real-time deformability cytometry: On-the-fly cell mechanical phenotyping. Nat. Methods
**2015**, 12, 199–202. [Google Scholar] [CrossRef] [PubMed] - Bruker AFM Probes, “MLCT-BIO.” [Online]. Available online: https://www.brukerafmprobes.com/p-3945-mlct-bio.aspx (accessed on 2 February 2021).
- Xia, Y.; Whitesides, G.M. Soft Lithography. Annu. Rev. Mater. Sci.
**1998**, 28, 153–184. [Google Scholar] [CrossRef] - Jötten, A.M. SURF Survival of Rosettes in Flow. 2021. Available online: https://github.com/anna-joe-0305/SURF (accessed on 14 November 2021).
- Jötten, A.M.; Moll, K.; Wahlgren, M.; Wixforth, A.; Westerhausen, C. Blood group and size dependent stability of P. falciparum infected red blood cell aggregates in capillaries. Biomicrofluidics
**2020**, 14, 024104. [Google Scholar] [CrossRef] - Liljequist, D.; Elfving, B.; Roaldsen, K.S. Intraclass correlation—A discussion and demonstration of basic features. PLoS ONE
**2019**, 14, e0219854. [Google Scholar] [CrossRef] [Green Version] - Mahaffy, R.E.; Shih, C.K.; MacKintosh, F.C.; Käs, J. Scanning Probe-Based Frequency-Dependent Microrheology of Polymer Gels and Biological Cells. Phys. Rev. Lett.
**2000**, 85, 880–883. [Google Scholar] [CrossRef] [Green Version] - Hayashi, K.; Iwata, M. Stiffness of cancer cells measured with an AFM indentation method. J. Mech. Behav. Biomed. Mater.
**2015**, 49, 105–111. [Google Scholar] [CrossRef] - Galajda, P.; Kelemen, L.; Végh, G.A. Micro- and nanotechnology for cell biophysics. Acta Biol. Szeged.
**2015**, 59, 303–321. [Google Scholar] - Li, Q.S.; Lee, G.Y.H.; Ong, C.N.; Lim, C.T. AFM indentation study of breast cancer cells. Biochem. Biophys. Res. Commun.
**2008**, 374, 609–613. [Google Scholar] [CrossRef] [PubMed] - Ren, J.; Huang, H.; Liu, Y.; Zheng, X.; Zou, Q. An atomic force microscope study revealed two mechanisms in the effect of anticancer drugs on rate-dependent Young’s modulus of human prostate cancer cells. PLoS ONE
**2015**, 10, e0126107. [Google Scholar] [CrossRef] [PubMed] - Pogoda, K.; Jaczewska, J.; Wiltowska-Zuber, J.; Klymenko, O.; Zuber, K.; Fornal, M.; Lekka, M. Depth-sensing analysis of cytoskeleton organization based on AFM data. Eur. Biophys. J.
**2012**, 41, 79–87. [Google Scholar] [CrossRef] [PubMed] - Mokbel, M.; Mokbel, D.; Mietke, A.; Träber, N.; Girardo, S.; Otto, O.; Guck, J.; Aland, S. Numerical Simulation of Real-Time Deformability Cytometry to Extract Cell Mechanical Properties. ACS Biomater. Sci. Eng.
**2017**, 3, 2962–2973. [Google Scholar] [CrossRef] [PubMed]

**Figure 1.**(

**A**) Micrograph of a cantilever without tip approaching an adherent A375 cell. (

**B**) Principle AFM with and without tip. (

**C**) Force–distance curve of an exemplary measurement with cantilever stiffness k = 0.03 N/m, force set point f = 2 nN, approaching speed v = 1 µm/s and approaching distance d = 5 µm. The pause between approach and retract is 1 s at constant force. (

**D**) Comparison of AFM force spectroscopy results of the Young’s modulus of A375 cells using a cantilever with tip and one without tip. The boxplot summarizes the results on about 50 cells with approximately ten force curves per cell of AFM measurements with and without tip. The cantilever without tip is 20 μm wide at the front, and the tip radius of the approximately 3 μm long tips used here is 20 nm.

**Figure 2.**AFM force spectroscopy mapping of a cell using a cantilever with tip. (

**A**) Map of half a cell, measurement started at the center of the cell. (

**B**) Typical force distance curve from the softer outer area, (

**C**) typical force distance curve from the central, stiffer part.

**Figure 3.**Young’ modulus E as a function of the loading rate normalized to the Young’s modulus from reference measurements with a loading rate a = 1.0 µm/s. The logarithmic fit function is E = 0.98 + 0.19 ln (a + 0.1).

**Figure 4.**(

**A**) Design of the microfluidic stenosis narrowing from diameter d

_{1}to diameter d

_{2}over the elongational length e. The cell deforms from a round shape with its original diameter d = 2r

_{0}to an ellipse with major axis L. (

**B**) Exemplary micrographs of A375 cells passing a standard elongation d = 7 µm stenosis. The area for the deformation analysis is highlighted in red.

**Figure 5.**(

**A**) Velocity, major and minor ellipse axes from one measurement as function of the position x. Mean and standard deviation from 2300 cell trajectories. (

**B**) Stress–strain diagram and linear fit to determine the Young’s modulus of one set of A375 cells. (

**C**) Resulting Young’s moduli from six independent measurements as shown in B, representing about 2500 cells each.

**Figure 6.**Cell deformation as a function of the x-position; the center of the stenosis is at x = 0. The shape of the cell is approximated by an ellipse. The eccentricity ε of an ellipse is the distance between the foci divided by the major axis length. The labelled ellipses illustrate the values. “Long” and “short” stenosis refers to stenoses with length e = 165 µm and e = 55 µm, respectively.

**Figure 7.**Comparison of AFM and microfluidic deformation analysis of the Young’s modulus of A375 cells. AFM: The result of AFM measurements with and without tip is the mean and standard deviation from measurements on 5–15 cells with $\approx $ 10 force curves per cell. Stenosis: Elasticity was also determined based on deformation under flow during the passage of a stenosis. The value here summarizes the six values from Figure 5, each of which relies on trajectories from approximately 2500 cells.

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**MDPI and ACS Style**

Jötten, A.M.; Neidinger, S.V.; Tietze, J.K.; Welzel, J.; Westerhausen, C.
Dynamic Effective Elasticity of Melanoma Cells under Shear and Elongational Flow Confirms Estimation from Force Spectroscopy. *Biophysica* **2021**, *1*, 445-457.
https://doi.org/10.3390/biophysica1040032

**AMA Style**

Jötten AM, Neidinger SV, Tietze JK, Welzel J, Westerhausen C.
Dynamic Effective Elasticity of Melanoma Cells under Shear and Elongational Flow Confirms Estimation from Force Spectroscopy. *Biophysica*. 2021; 1(4):445-457.
https://doi.org/10.3390/biophysica1040032

**Chicago/Turabian Style**

Jötten, Anna Martina, Simon V. Neidinger, Julia K. Tietze, Julia Welzel, and Christoph Westerhausen.
2021. "Dynamic Effective Elasticity of Melanoma Cells under Shear and Elongational Flow Confirms Estimation from Force Spectroscopy" *Biophysica* 1, no. 4: 445-457.
https://doi.org/10.3390/biophysica1040032