Atomic Force Microscopy Reveals Integrin β1 Dynamics During Human Adipose‑Derived Stem Cell Chondrogenesis

Abstract

Integrin β1 orchestrates key processes such as differentiation, migration, and tissue repair. Using atomic force microscopy (AFM), we quantified the interaction between integrin β1 ligands and CD29 receptors on human adipose‑derived stem cells (hADSCs) throughout early chondrogenic differentiation. AFM imaging showed a transition from elongated, spindle‑shaped cells to polygonal cells with reduced length/width ratios and increased surface roughness. Over 1,200 force–distance curves collected at 0, 6, and 12 days revealed rising rupture forces (61.8 ± 22.2 pN → 67.2 ± 22.0 pN) and binding probabilities (19.6 % → 33.4 %). These data demonstrate that AFM can track integrin β1‑CD29 engagement, cellular ultrastructure, and morphology as reliable markers of chondrogenic progression.

Background

Osteoarthritis (OA) erodes articular cartilage, a tissue that is avascular and has limited self‑repair. Stem‑cell‑based tissue engineering holds promise for restoring cartilage function, yet the precise signaling events guiding chondrogenesis remain unclear. Integrins, particularly integrin β1 (CD29), mediate cell–matrix interactions and activate pathways such as Wnt/β‑catenin, PI3K, and mTOR that govern stem‑cell fate. We focused on integrin β1 because of its pivotal role in cartilage biology and its interaction with diverse α subunits.

AFM, coupled with single‑molecule force spectroscopy (SMFS), offers nanometer‑resolution imaging of living cells and direct measurement of ligand–receptor forces. By functionalizing AFM tips with integrin β1 antibodies, we probed binding events on hADSCs during chondrogenic induction, providing kinetic and structural insights unavailable by conventional assays.

Methods

Cell Culture and Reagents

hADSCs were isolated from subcutaneous fat of three donors (mean age 20 y) under IRB‑approved protocols. Cells were expanded in low‑glucose DMEM supplemented with 10 % heat‑inactivated FBS, antibiotics, sodium pyruvate, and L‑glutamine, and maintained at 37 °C, 5 % CO₂.

In Vitro Differentiation

For chondrogenesis, passage 4–8 cells were seeded at 2 × 10⁵ cells per 10 mL and cultured in chondrogenic medium containing 1 % FBS, ITS, 10 ng/mL TGF‑β1, 100 ng/mL IGF‑1, 10⁻⁷ M dexamethasone, and 50 µg/mL ascorbic acid. Medium changes occurred every 2 days. Cartilage matrix deposition was confirmed by Alcian blue and toluidine blue staining.

Osteogenic and adipogenic differentiation followed standard protocols with dexamethasone, ascorbic acid, β‑glycerol phosphate, IBMX, hydrocortisone, and indomethacin, assessed by alizarin red and Oil Red O staining, respectively.

Flow Cytometry

Cells were labeled with antibodies against CD34, CD44, CD45, CD73, CD90, CD106, HLA‑DR, and CD105 and analyzed to confirm mesenchymal phenotype.

Immunoblotting and Immunofluorescence

Protein extracts were probed for β‑catenin, integrin β1, collagen II, GSK‑3β, SOX, and β‑actin. Immunofluorescence visualized integrin β1 distribution and F‑actin organization using Alexa Fluor 488 and phalloidin‑Alexa Fluor 573, respectively.

AFM Tip Functionalization

Silicon nitride tips were cleaned, silanized with 1 % 3‑APTES, cross‑linked with glutaraldehyde, and incubated overnight with 1 mg/mL anti‑integrin β1 antibody to generate ligand‑functionalized probes.

AFM Measurements

Cells were fixed with 4 % paraformaldehyde and imaged in PBS. Morphology and surface roughness (Ra, Rq) were quantified from >15 10 µm² images per time point. SMFS was performed in approach–retract mode at 500 nm/s, collecting >400 force curves per experiment (≈ 1,200 total per time point). Binding events were identified by rupture forces >800 pN and a force threshold calibrated to 0.058 ± 0.006 N/m.

RT‑qPCR

RNA was extracted, reverse‑transcribed, and quantified for integrin β1 and GAPDH using SYBR Green chemistry. Relative expression was calculated by the 2^–ΔΔCT method.

Statistics

Experiments were performed in triplicate. Data are presented as mean ± SD. Comparisons used t‑tests or one‑way ANOVA with Bonferroni post‑tests; p < 0.05 was considered significant.

Results and Discussion

hADSC Characterization

Flow cytometry confirmed the mesenchymal phenotype (CD73⁺, CD90⁺, CD105⁺, CD34⁻, CD45⁻). MTT assays demonstrated robust proliferation across passages.

Morphological and Ultrastructural Shifts During Chondrogenesis

AFM revealed a clear morphological shift from spindle‑like to polygonal cells, with a significant reduction in length/width ratio at days 6 and 12 (p < 0.01). Surface roughness parameters Ra and Rq increased markedly, indicating enhanced membrane heterogeneity likely driven by ECM remodeling.

Cytoskeletal Remodeling

Confocal imaging showed F‑actin reorganization from a longitudinal array to a radial, peripherally‑localized pattern by day 12, reflecting cytoskeletal remodeling associated with chondrogenic commitment.

Integrin β1–CD29 Binding Dynamics

SMFS with integrin β1‑functionalized tips yielded increasing rupture forces and binding probabilities over time (0 % → 33.4 % at 12 days). Blocking experiments with free anti‑integrin β1 antibody reduced force curves by ~90 %, confirming specificity.

Integrin β1 Expression and Signaling

Western blot and qRT‑PCR demonstrated up‑regulation of integrin β1, collagen II, and components of the β‑catenin/SOX pathway during chondrogenesis. Elevated β‑catenin, SOX, and GSK‑3β levels suggest that integrin β1 activates Wnt/β‑catenin signaling to drive cartilage matrix synthesis.

Implications and Limitations

AFM provides a non‑destructive, single‑cell approach to monitor integrin dynamics, morphology, and membrane roughness as real‑time indicators of chondrogenic differentiation. Future studies should investigate integrin conformation and delineate the precise molecular links between integrin β1 and β‑catenin/SOX signaling.

Conclusions

We established AFM as a powerful tool to track integrin β1–CD29 interactions and nanostructural changes during hADSC chondrogenesis. The technique offers kinetic and structural insights that can inform stem‑cell‑based cartilage repair strategies.