Chromatin decondensation and nuclear softening accompany Nanog downregulation in embryonic stem cells

Abstract

The interplay between epigenetic modification and chromatin compaction is implicated in the regulation of gene expression, and it comprises one of the most fascinating frontiers in cell biology. Although a complete picture is still lacking, it is generally accepted that the differentiation of embryonic stem (ES) cells is accompanied by a selective condensation into heterochromatin with concomitant gene silencing, leaving access only to lineage-specific genes in the euchromatin. ES cells have been reported to have less condensed chromatin, as they are capable of differentiating into any cell type. However, pluripotency itself-even prior to differentiation-is a split state comprising a naïve state and a state in which ES cells prime for differentiation. Here, we show that naïve ES cells decondense their chromatin in the course of downregulating the pluripotency marker Nanog before they initiate lineage commitment. We used fluorescence recovery after photobleaching, and histone modification analysis paired with a novel, to our knowledge, optical stretching method, to show that ES cells in the naïve state have a significantly stiffer nucleus that is coupled to a globally more condensed chromatin state. We link this biophysical phenotype to coinciding epigenetic differences, including histone methylation, and show a strong correlation of chromatin condensation and nuclear stiffness with the expression of Nanog. Besides having implications for transcriptional regulation and embryonic cell sorting and suggesting a putative mechanosensing mechanism, the physical differences point to a system-level regulatory role of chromatin in maintaining pluripotency in embryonic development.

Figure 1

(A) Schematic representation of forces acting on cells and nuclei during optical stretching. Cells are trapped and stretched between two counterpropagating laser beams. Refractive index changes between the surrounding medium and the cell lead to a momentum transfer from light to the cell surface. The resulting forces are schematically represented by black arrows (left). In addition to forces acting on the cell surface, refractive index changes between the cytoplasm and the nucleus lead to a deformation of the nucleus (blue arrows, right). Optical stretching can be used to probe nuclear mechanics within living cells, where nuclei are visualized with DNA-specific dyes (Hoechst). (B) Representative examples of a cell and a nucleus that have been trapped (0.4 W total laser power) and stretched (2.2 W) in optical-stretching experiments. Scale bar (same for cell and nucleus), 1 μm. Edge detection was used to measure the relative deformation along the laser axis: Strain = (L1 − L0)/L0.

Figure 2

Optical stretching experiments show that LN cells, as well as LN nuclei, are more deformable than HN cells and HN nuclei, respectively. (A) Strain-versus-time plot for HN and LN cells. The stretching period is indicated by the red bar. (B) Cellular peak strains of HN and LN cells in OS experiments. The secondary y axis gives the values for cell compliance (see Methods). (C) TNGA cells were treated with MgCl₂ + CaCl₂ to condense, or TSA to decondense, chromatin y axis, and cellular peak strains were compared to an untreated control population. (D) Cells were treated with CytoD or Noco to disrupt actin or microtubule filaments, respectively. Note that the relative differences between HN and LN cells remain unchanged upon cytoskeleton disruption. (E) Strain-versus-time plot for the deformation of HN and LN nuclei stretched within intact cells. (F) Nuclear peak strains for HN and LN populations. (G) Treatment of TNGA cells with chromatin-modifying agents led to changes in nuclear deformability relative to an untreated control population (AZA decondenses chromatin). These changes correspond well to differences in whole-cell deformability (compare to C). Data in plots A and E are given as the mean ± SE from n individual measurements. Data in all bar diagrams represent the mean ± SD of N independent experiments unless stated otherwise. Statistical significance is indicated by asterisks: ^∗p ≤ 0.05; ^∗∗p ≤ 0.01; ^∗∗∗p ≤ 0.001.

Figure 3

Confocal imaging and quantification of heterochromatin markers in TNGA cells. (A) TNGA cells (HN, GFP-positive; LN, GFP-negative) were treated with TSA to decondense chromatin and stained for HP1α, a marker for heterochromatic foci (middle column). The fractal dimension, D, was calculated for individual nuclei; examples are highlighted and D value is indicated. (B) TNGA cells stained for the heterochromatin marker H3K27Me3. (C) Fractal dimension was quantified for HP1α and H3K27Me3 distributions and compared between HN and LN cells. Note that a higher fractal dimension corresponds to a more homogeneous distribution pattern. (D) Quantitative results of the experiment described in A. (E) HN cells show a higher staining intensity for both H3K27Me3 and Oct4 than do LN cells, indicating a higher expression of both heterochromatin and pluripotency markers. Scale bar is 5 μM.

Figure 4

FRAP experiments for labeled histone proteins. (A) Histone H2B was fluorescently labeled with RFP and a bleach pulse was applied to a circular area. (B) Fluorescence recovery in the bleached area was recorded for 600 s for HN and LN cells. Data were fit to an exponential decay function, f(t) = 1 − exp(−t/τ), and time constant, τ, is reported in the figure key. (C) At the end of the observation period, LN cells show a significantly higher mean recovery than HN cells (inset; mean ± SD from four independent experiments). (D) HN cells treated with the histone deacetylase inhibitor TSA to decondense chromatin show fluorescence recovery kinetics similar to those observed for LN cells (mean ± SE from one representative experiment).