DNA damage regulates the mobility of Brca2 within the nucleoplasm of living cells

Edited by Stephen C. Kowalczykowski, University of California, Davis, CA, and approved October 26, 2010 (received for review July 13, 2010)
November 22, 2010
107 (50) 21937-21942


How the biochemical reactions that lead to the repair of DNA damage are controlled by the diffusion and availability of protein reactants within the nucleoplasm is poorly understood. Here, we use gene targeting to replace Brca2 (a cancer suppressor protein essential for DNA repair) with a functional enhanced green fluorescent protein (EGFP)-tagged form, followed by fluorescence correlation spectroscopy to measure Brca2-EGFP diffusion in the nucleoplasm of living cells exposed to DNA breakage. Before damage, nucleoplasmic Brca2 molecules exhibit complex states of mobility, with long dwell times within a sub-fL observation volume, indicative of restricted motion. DNA damage significantly enhances the mobility of Brca2 molecules in the S/G2 phases of the cell cycle, via signaling through damage-activated protein kinases. Brca2 mobilization is accompanied by increased binding within the nucleoplasm to its cargo, the Rad51 recombinase, measured by fluorescence cross-correlation spectroscopy. Together, these results suggest that DNA breakage triggers the redistribution of soluble nucleoplasmic Brca2 molecules from a state of restricted diffusion, into a mobile fraction available for Rad51 binding. Our findings identify signal-regulated changes in nucleoplasmic protein diffusion as a means to control biochemical reactions in the cell nucleus.
The error-free repair of DNA breaks in the cell nucleus is essential for genome integrity (1). Signals emanating from broken DNA trigger the recruitment of diffusing proteins from the nucleoplasm into microscopically visible “foci” (1) at the positionally stable damage sites (2), where repair reactions are carried out. Whether these events involve signal-initiated changes in the nucleoplasmic protein pool is unclear. The diffusion and availability of protein reactants in the nucleoplasm is influenced by molecular crowding and specific intermolecular interactions that incorporate them into macromolecular complexes of varying size (3). However, changes in nucleoplasmic protein diffusion are inaccessible to traditional biochemical methods, and so their involvement in DNA repair (or other site-specific DNA transactions) remains largely unexplored.
To address this problem, we have studied DNA double-strand break (DSB) repair by homologous recombination (HR), a fundamental biological process mediated in vertebrate cells by the assembly in repair foci of the cancer suppressor protein, Brca2, with its cargo, the recombination enzyme Rad51 (4). We have combined somatic gene targeting with fluorescence correlation spectroscopy (FCS) [a method successfully applied to the analysis of fluorescent particles in solution (5) or within cells (6)] to directly measure the diffusion of EGFP-tagged Brca2 expressed natively in the nucleoplasm of living cells. Here, we show that the signals triggered by DNA breakage initiate significant changes in nucleoplasmic Brca2 diffusion, suggestive of release from a state of restricted mobility, accompanied by an increase in its interaction with Rad51 in the same compartment detected by fluorescence cross-correlation spectroscopy (FCCS) (7). Our findings provide fresh insight into the control of HR reactions mediated by Brca2 and Rad51 and demonstrate signal-initiated changes in the diffusion of nucleoplasmic proteins during a biological process.


Replacement of Endogenous Brca2 with a Functional GFP-Tagged Form by Gene Targeting in DT40 Cells.

We used gene targeting to replace the endogenous Brca2 protein with a functional, fluorescently labeled form. This procedure overcomes the limitations of conventional approaches where fluorophore-tagged proteins are heterologously expressed in vertebrate cells for spectroscopic analysis (8), retaining the endogenous, untagged protein. We deleted one Brca2 allele in the avian DT40 cell line (9), a widely used experimental model for HR (10), and “knocked in” the gene encoding enhanced green fluorescent protein (EGFP) to the 3′ end of the coding sequence of the second allele, yielding a final genotype of Brca2.EGFP/− (Fig. 1 AC, SI Methods, and Table S1). In cell extracts, anti-EGFP reacts with only a single species in Western blots corresponding to Brca2-EGFP (Fig. S1A, Fig. 1D); alternative EGFP species that might confound fluorescence measurements are absent. An antibody against Gallus gallus (gg) Brca2 detects a band with slightly retarded mobility relative to untagged Brca2, but not the untagged protein (Fig. 1D). Nuclear Brca2-EGFP at endogenous levels can be distinguished from the predominantly cytoplasmic autofluorescence background, but the typical fluorescence intensities are insufficient for image-based analyses (Fig. 1E, Fig. S1 BD). Staining with an anti-GFP antibody in fixed cells confirms that Brca2-EGFP is mainly nuclear and expressed at varying levels in the asynchronous cell population, as expected (Fig. S1 E and F). Several lines of evidence confirm that Brca2-EGFP functionally replaces endogenous Brca2. First, Brca2-EGFP forms foci after DNA damage, which colocalize with Rad51 (Fig. S2A). Second, Brca2.EGFP/− cells are as efficient as parental cells in supporting Rad51 focus formation after DNA damage (Fig. S2B). Finally, Brca2.EGFP/− cells have no measurable deficit in DNA repair by HR compared with control parental cells. They are neither hypersensitive to the DNA cross-linking agent, mitomycin C (MMC) (Fig. S2C), nor defective in sister chromatid exchange (SCE), a direct measure of HR (10), before or after exposure to MMC (Fig. S2 D and E). Thus, we have created a unique experimental model to study changes in Brca2 diffusion during physiological reactions, using quantitative fluorescence imaging.
Fig. 1.
Construction and visualization of an endogenously tagged fluorescent Brca2 protein. (A) Generation of the Brca2.EGFP/− DT40 cell line. One Brca2 allele was excised to create Brca2/− heterozygote cells (11). EGFP was then targeted to the 3′ end of the wild-type allele to generate Brca2.EGFP/− cells. (B) The targeting constructs. Primers For-B2GFP and Rev-B2GFP (arrows) amplify by PCR (C) a 4.5-kb band only from the EGFP-targeted allele in Brca2.EGFP/− cells (lane 2) or a 0.6-kb band from the wild-type allele in Brca2/− cells (lane 3). (D) Brca2.EGFP/− cells express a full-length Brca2-EGFP fusion protein (black arrow) detected by anti-GFP or anti-Gg Brca2 as a band with slightly retarded gel mobility compared with Brca2 (gray arrow). (E) Brca2.EGFP localization in a living cell imaged with a laser scanning microscope (10-μm optical slice, averaged ×16).

Diffusion of Soluble Nucleoplasmic Brca2-EGFP Measured by Fluorescence Correlation Spectroscopy.

We used FCS to dissect the diffusion of molecular complexes containing Brca2-EGFP within the nucleus of living cells. FCS measures (5) the temporal fluctuations in fluorescence arising when fluorophores diffuse through a small confocal volume (Fig. S3 A and B). Their correlation over time is described by the autocorrelation function (ACF) or G(τ) (Fig. S3A; Methods). The ACF depends on the average concentration of fluorescent species and their dwell time (τD) within the observation volume, permitting an estimate of the diffusion coefficient and number of fluorescently tagged molecules (for further information, see ref. 8, SI Methods). Two features of FCS make the technique ideal to study the diffusion of Brca2. First, FCS is particularly suited to fluorescent molecules like endogenous Brca2-EGFP expressed at low concentrations. Second, FCS preferentially reports the properties of soluble Brca2 molecules in the nucleoplasm (11), a fraction that has not been previously characterized.
Although the observation volume for our instrument (∼0.25 fL) is not small enough to entirely exclude chromatin domains or small subnuclear structures, any fluorescent molecules stably bound to these domains or structures are photobleached very rapidly when observation commences (e.g., the fluorescence trace in Fig. S3C). However, initial photobleaching is absent from the majority of FCS traces (e.g., Fig. S3B) and is likely related to the occasional presence of foci of stable molecules close to the FCS measurement site (Fig. S3 legend and Methods). This result is consistent with the observation that in the few cells expressing sufficient Brca2-EGFP to enable fluorescence recovery after photobleaching (FRAP), >60% of nuclear Brca2 is mobile (Fig. S4 A and B). Nevertheless, most cells are not amenable to FRAP analysis because the limited fluorescence signal emitted by the low concentration of endogenously expressed Brca2-EGFP was too small to permit robust quantification (Fig. S4C). We therefore used FCS to focus on the analysis of mobile Brca2-EGFP molecules, excluding the immobile fraction in the initial segment of the fluorescence traces. Mobile Brca2-EGFP molecules are collectively termed the “soluble nucleoplasmic pool.” This fraction includes molecules that undergo transient binding for times less than the period required for fluorophore bleaching during observation (8). The fluorescence of diffusing Brca2-EGFP molecules in the nucleoplasm can be clearly distinguished from the autofluorescent background (Fig. S3D), showing higher signals and autocorrelation compared with measurements performed on parental lines.

Modeling of Brca2-EGFP FCS Measurements.

The mobility of fluorescent molecules in solution can be characterized by fitting the ACF to appropriate models (12). The simplest diffusion process that can be described by FCS measurements is random 3D diffusion (12, 13) for a single component (Methods). This simple model fits the ACF of a biologically inert tracer, EGFP (Fig. 2A), but not the complex diffusion pattern of Brca2-EGFP (Fig. 2 A–C).
Fig. 2.
A pool of nucleoplasmic Brca2-EGFP with restricted mobility. (A) Representative ACFs for Brca2-EGFP (blue squares) or free EGFP (open black circles), each fitted to a single-component 3D diffusion model (red lines). (B) Distribution of amplitude-normalized ACFs from 70 Brca2.EGFP/− cells. Each curve is the average of ten 5-s readings in a single cell. The residuals from fitting to three different diffusion models are shown in C. Error bars represent the SEM. (D) Distribution of the fast-diffusing Brca2-EGFP component described by the two-component model in 70 Brca2.EGFP/− cells. (E) Comparison of the measured dwell time (τD) of the fast- and slow-diffusing Brca2-EGFP components (gray bars) in 70 Brca2.EGFP/− cells with free EGFP (black bars; n = 30). The population average was used to calculate the apparent D values noted in the text.
Nucleoplasmic Brca2-EGFP diffusion may be altered by its participation in protein complexes or hindered by molecular crowding, transient intermolecular interactions, or obstructions to free diffusion created by chromatin and other immobile structures (14). We therefore applied two typical models: the two-component 3D diffusion model, incorporating a second diffusing species (12, 15), or alternatively a model accounting for anomalous or “subdiffusive” processes (16) (Methods). Subdiffusive processes represent situations where a particle in a random walk spends a higher-than-average time at each point before moving on (reflected mathematically as the anomaly parameter), which may occur within the nucleoplasm (14). Indeed, the goodness of fit (the residual sum of squares) for FCS measurements from a population of Brca2.EGFP/− cells was improved relative to a single-component model by including either anomalous diffusion or a second diffusive species (Fig. S5 A and B).
In the first case, fitting of the ACF yields an anomaly parameter, α = 0.67 ± 0.01, similar to that obtained for other nuclear proteins (14), and a dwell time for anomalous diffusion of τDα = 3.9 ± 0.2 ms (Fig. S5 C and D). However, the standardized residuals for the anomalous diffusion fit show systematic deviations that are not present when the same data are analyzed with a two-component model (Fig. 2C). Thus, a single anomalously diffusing component is insufficient to describe Brca2-EGFP behavior. Indeed, a multicomponent model is more biologically intuitive for DNA repair proteins like Brca2, known to form soluble complexes involving different binding partners (17). We used ATP depletion to dissect the contribution of binding events to Brca2-EGFP mobility (SI Discussion). ATP depletion decreases the average time of anomalous diffusion for Brca2-EGFP, whereas the anomaly parameter itself is unaffected. This result suggests that energy-dependent binding events occur within the diffusing pool, regardless of the model chosen (Fig. S6 and SI Discussion). Together, these considerations indicate that in our biological system a two-component 3D-diffusion model is the best minimal choice for data analysis.

A Pool of Nucleoplasmic Brca2-EGFP with Restricted Mobility.

The two-component 3D-diffusion model identifies a faster-diffusing Brca2-EGFP component accounting for ∼50% (47.9 ± 1.2%, mean ± SE, n = 70 cells) of the total pool (Fig. 2D), which has an average diffusion coefficient, D = 14.8 ± 1.0 μm2/s (Fig. 2E). This is ∼2.1-fold lower than that of free EGFP (D = 30 ± 2 μm2/s, n = 22). Although the diffusion coefficient of proteins in vivo cannot be reliably predicted (13, 14), the expected difference in mass between EGFP (27 kDa) and Brca2-EGFP (411 kDa) suggests that the fast component can be interpreted as free, or minimally bound Brca2-EGFP protein (SI Discussion). The remaining ∼50% of the diffusing Brca2-EGFP pool is represented by a significantly slower component with an average dwell time of τD = 22.1 ± 1.4 ms (n = 70) (Fig. 2E). The slow component can be interpreted as either the diffusion of gigantic complexes (∼8 GDa) of Brca2-EGFP (D = 0.45 ± 0.03 μm2/s) or, alternatively, transient immobilization of the protein due to binding events (koff = 1/τD ∼ 45 s−1) and likely represents a combination of complex formation and transient binding events (Fig. S7 A and B). Satisfactory fitting to the two-component model does not exclude the presence of anomalous diffusion within the nucleus; further dissection is not currently possible by FCS alone. Importantly, however, both models suggest that a significant fraction of nucleoplasmic Brca2-EGFP molecules reside within pools of restricted mobility (Figs. S6 and S7 A and B and SI Discussion).
Fit estimates from the analysis of the FCS measurements show significant scatter between cells, which could represent biological differences between cells endogenously expressing Brca2-EGFP under native control. However, a similar distribution of values is also observed among multiple readings taken within a single cell, suggesting that our sampling accurately represents the average behavior of functional Brca2-EGFP molecules (Fig. S7 C–F).

DNA Damage Mobilizes Brca2-EGFP, but Not Inert Fluorescent Tracers.

DNA damage induces significant changes in the diffusion of Brca2-EGFP, suggestive of its release from pools with restricted mobility. This conclusion is apparent from a comparison of the population mean ± SE of amplitude-normalized ACF curves from cells before, and 1–3 h after (Fig. 3A), exposure to 10 Gy of ionizing radiation (IR). The leftward shift reflects a shorter average dwell time for fluorescent molecules in the observation volume, demonstrating an overall increase in the diffusional mobility of Brca2-EGFP after DNA damage. In contrast, two sets of measurements 4 h apart on populations of undamaged cells are identical (Fig. 3B). The observed ACF change after DNA damage (noting the logarithmic scale used in Fig. 3A) is consistent with a substantial reorganization of Brca2-EGFP–containing macromolecular complexes in vivo. This change is unlikely to be caused by the mobilization of chromatin-bound Brca2 because prior studies using classical biochemical fractionation show that the overall amount of chromatin-bound Brca2 increases rather than decreases after DNA damage (18). Instead, the statistically significant increase we observe in the fraction of Brca2-EGFP molecules corresponding to the fast-diffusing component (Fig. 3 C and D, t test, P < 0.01) suggests a reorganization of Brca2-EGFP–containing macromolecular complexes visible under FCS into simpler forms with higher mobility.
Fig. 3.
DNA damage increases the mobility of Brca2-EGFP. (A) Mean amplitude-normalized ACF curves for Brca2-EGFP before (black) or 3 h after (red) exposure of Brca2.EGFP/− cells to 10 Gy IR; note the leftward shift of the red curve. Each data point is the average ± SEM of readings from 50 cells. Insets show Brca2-EGFP localization. (B) Mean amplitude-normalized ACF curves ± SEM for Brca2-EGFP from undamaged Brca2.EGFP/− cells at similar time points. (C) Per-cell values for the amount of fast-diffusing Brca2-EGFP in a two-component fit (an index of mobility derived from the ACF). Each circle represents the average from a single cell, and the red line shows the population mean ± SEM. DNA damage induces statistically significant changes (t test, P < 0.01, n ∼ 50 cells each). (D) Dot plot of the same dataset analyzed by globally linking the τD values, which shows a similar difference in fast-diffusing Brca2-EGFP. Numbers in parentheses show the τD (in μs) for the fast and slow Brca2-EGFP components from the linked fit. Mean amplitude-normalized ACF curves ± SEM for free EGFP (E, n = 20 cells each), for 10 kDa TMR-dextran (F, n = 25), and for Cy3-Oligo(dT) (G, n = 25) are shown before (black) and 1–3 h after (red) 10 Gy IR.
Damage-induced mobilization of Brca2-EGFP could arise either directly, from alterations in Brca2-EGFP's physical properties, or indirectly, through alterations in nuclear viscosity (SI Methods). To discriminate between these possibilities, we introduced inert molecular tracers to report changes in the nuclear microenvironment that might be induced by DNA damage. No detectable alterations were found. There is no shift in the mean ACF for free EGFP (Fig. 3D), a globular, compact molecule; for a 10-kDa TMR-dextran polymer that exhibits a greater hydrodynamic radius (SI Methods, Fig. 3E); or for Cy3-labeled oligo(dT) (Fig. 3F), a tracer incorporated into massive ribonucleoprotein complexes of the order of Brca2-EGFP in molecular mass (19). Thus, DNA damage neither triggers global alterations in nuclear viscosity nor grossly alters the shape of the observation volume within the cell. This result confirms that damage-induced Brca2-EGFP mobilization arises from specific changes in its physical properties. Collectively (and irrespective of the model chosen for FCS analysis), our experiments show that DNA damage changes nucleoplasmic Brca2 molecules, suggestive of redistribution from restricted mobility to a more mobile state.

Brca2-EGFP Mobilization Coincides with Active HR.

Several observations suggest that damage-induced Brca2-EGFP mobilization correlates with active HR. First, the altered mobility of Brca2-EGFP increases over time following IR damage, peaking 3–4 h afterward (Fig. 4A), similar to the time course for the assembly of HR foci at DSBs (20). Second, Brca2-EGFP mobilization does not occur if cells are exposed to IR during the G1 phase of the cell cycle (Fig. 4B), a period when HR foci do not form, and HR may be inactive (21). By contrast, there is a pronounced mobilization in cells exposed to IR during the S and G2 phases (Fig. 4C), corresponding to the period when HR is most active. Finally, the clastogenic agents VP16 and MMC, which generate DNA lesions that induce HR foci formation and engage the HR repair machinery (22), also trigger the mobilization of Brca2-EGFP (Fig. 4 D and E, respectively). These findings suggest that Brca2-EGFP mobilization is a biological response to DNA breakage that coincides with HR activity during the cell cycle.
Fig. 4.
Brca2-EGFP mobilization coincides with active HR. (A) Average amount ± SEM of fast-diffusing Brca2-EGFP in a two-component fit from 15 cells exposed to 10 Gy IR (binned at 1-h intervals). Time points >6 h are not shown due to increased cell death. (B) Mean amplitude-normalized ACF curves ± SEM for Brca2-EGFP from Brca2.EGFP/− cells arrested in G1 using mimosine before (black) or 2 h after (red) exposure to 10 Gy IR. Inset shows DNA content measured by propidium iodide staining and flow cytometry in asynchronous (gray), G1-arrested (black), or G1-arrested and IR-exposed (red) cells. A similar experiment is depicted in C, except that the cells are synchronized in S-G2 after release from mimosine arrest. (D) Percentage of fast-diffusing Brca2-EGFP in two-component fits from Brca2.EGFP/− cells before and after exposure to 20 μM Etoposide (VP16) for 2 h (P < 0.01, n = 20 each). (E) A similar plot for G1-synchronized cells released into 100 ng/mL MMC and followed for 6 h (P < 0.01, pre- vs. post-MMC 2 h and 4 h; n ∼ 50 each). Experimental design is depicted below the data. (F) A similar plot for cells before or 3 h after 10 Gy IR, with or without preexposure to 1 μM wortmannin for 30 min (P < 0.01, n ∼ 25 cells each). Each data point represents the average of 10 measurements from within a cell, with error bars at the 95% confidence interval. Wortmannin also inhibits γH2AX formation after 10 Gy IR, with β-actin serving as a loading control.

Brca2-EGFP Mobilization Is Triggered by Damage-Activated Protein Kinase Activity.

Protein kinases of the phosphoinositide-kinase–like (PIK) family, like ATM, ATR, and DNA-PK, signal DSBs to initiate repair by HR. Preexposure of cells to wortmannin (a potent inhibitor of these kinases) (23) inhibits H2AX phosphorylation, a known PIK-dependent response to DSBs. Strikingly, it also suppresses Brca2-EGFP mobilization after exposure to IR (Fig. 4F), suggesting that this change is initiated by DSB-activated protein kinase activity.
Excessive DNA damage may trigger apoptosis. However, apoptotic cells marked by propidium iodide incorporation after exposure to 10 Gy IR (Fig. S8A) show no autocorrelation for Brca2-EGFP (Fig. S8B), suggesting they do not contribute to the damage-induced mobilization seen by FCS. Moreover, irradiation of asynchronous cells with a lower dose of IR, 5 Gy, induced a moderate but statistically significant mobilization of Brca2-EGFP (Fig. S8C). When specifically analyzing cells synchronized in the S-G2 phase, where Brca2 activity is maximal, we find that these cells exposed to 5 Gy IR exhibit a response similar to that of 10 Gy irradiated S-G2 cells, although the extent of apoptosis was considerably lower (Fig. S8 D–G). Together, these results argue that damage-induced Brca2-EGFP mobilization is unrelated to apoptosis. This conclusion is also supported by the observation that wortmannin, which enhances IR-induced cell death (24), nonetheless suppresses mobilization.

Brca2-EGFP Mobilization Is Accompanied by an Increase in Binding to Rad51.

Collectively, our findings provide several lines of evidence that DNA damage induces the mobilization of Brca2-EGFP from a slowly diffusing state in the nucleoplasm. To test the functional significance of this biological response, we used FCCS (Fig. S9A) with the well-characterized EGFP/mCherry fluorophore pair (25) to examine the interaction of Brca2-EGFP with its cargo, Rad51, in the nucleoplasm before and after DNA damage. When two populations of spectrally distinct fluorophores diffuse together within the observation volume, the amplitude of their cross-correlation is directly proportional to the number of interacting molecules (26). We generated a Brca2.EGFP gene-targeted cell line coexpressing an mCherry-Rad51 fusion protein (which we have previously shown to functionally replace endogenous Rad51) (27). mCherry-Rad51 in Brca2.EGFP cells was stably expressed at levels similar to those of the endogenous protein, (Fig. S9B) and colocalized with Brca2-EGFP in damage-induced HR foci (Fig. S9C). FCCS analysis in these cells (corrected for possible errors due to background autofluorescence, spectral cross-talk, or chromatic aberrations as previously described in ref. 26) (SI Methods) showed a significant cross-correlation signal, in contrast to measurements on Brca2-EGFP and free mCherry (Fig. 5A). To quantify the extent of interaction between Brca2-EGFP and mCherry-Rad51 in its simplest case (SI Methods), we calculated the ratio of the cross-correlation amplitude to the lower autocorrelation amplitude (Gx/Gr, because in this case the Gr < Gg, where Gx is the cross-correlation amplitude, and Gg and Gr are the autocorrelation amplitudes of EGFP- and mCherry-tagged proteins, respectively) (26). Interestingly, we observe a significant increase in the Gx/Gr ratio (P < 0.01; t test, n = 40 each) after DNA damage (Fig. 5B). This observation reflects an increase in the extent of interaction between Brca2-EGFP and mCherry-Rad51 within the diffusible pool of nucleoplasmic proteins, accompanying damage-induced Brca2-EGFP mobilization in this pool. Thus, our results suggest that DNA damage triggers the redistribution of soluble nucleoplasmic Brca2 molecules from a state of restricted diffusion into a mobile fraction available for Rad51 binding.
Fig. 5.
Brca2-EGFP mobilization is accompanied by increased association with mCherry-Rad51 in the nucleoplasm. (A) Cross-correlation between Brca2-EGFP and mCherry-Rad51 (black) vs. Brca2-EGFP and free mCherry (red). (B) Cross-correlation index for Brca2-EGFP and mCherry-Rad51, defined as the ratio of cross-correlation to the lower autocorrelation value (Gx/Gr), plotted before (black) and after (blue) exposure to 10 Gy IR (P < 0.01, t test, n = 40 each). (C) Signal-initiated changes in nucleoplasmic Brca2 diffusion: a hypothetical model. In undamaged cells (Left), soluble nucleoplasmic Brca2 molecules (blue-crossed yellow circles) visible under FCS can be represented as a fast-diffusing component (Upper Left) or a slow-diffusing fraction showing restricted mobility (Upper Right). Immobile Brca2 (e.g., stably bound to DNA) is invisible under FCS (Lower, gray background). DNA damage (Right) mobilizes Brca2 from dynamic sequestration in the nucleoplasm, increasing its availability for binding to Rad51 (red triangles, Upper Left and Right). These changes could enhance the delivery of Brca2/Rad51 complexes to positionally stable DSBs (Lower).


Here, we have combined gene targeting with FCS to study the diffusion of functional, EGFP-tagged Brca2 molecules expressed natively within the nucleoplasm of living cells. In the absence of endogenous untagged Brca2, FCS measurements account for all functional mobile pools of the protein, enabling correlation with the biological process under study. Our work highlights the use of gene targeting in DT40 cells to study the diffusion of fluorescent proteins (28) and provides a template for future FCS studies on DNA transactions in the cell nucleus.
We find that Brca2 molecules exist in complex states of mobility in the nucleoplasm. Their behavior can be interpreted biologically to imply the existence of transient or stable complexes (using the two-component model for 3D diffusion) (8) or as an obstruction to free diffusion through crowding, compartmentalization, or transient binding events (using the anomalous diffusion model) (16). In either case, a significant fraction of Brca2 within the nucleoplasm of living cells resides in a pool with restricted mobility (SI Discussion). Strikingly, DNA damage significantly enhances Brca2 mobility, without evidence of a global change in nucleoplasmic diffusion, implying that it triggers the redistribution of Brca2 from slow-moving forms into a more mobile state (Fig. 5C). Several lines of evidence indicate that the observed changes in Brca2-EGFP diffusion are biologically relevant. They are not only statistically significant across several independent observations, but also correlate well with the known biological activity of Brca2. Moreover, these changes coincide with active HR and are dependent on signaling through DNA damage-activated kinases of the PIK family, characteristic of a specific, signal-initiated cellular response to DNA breakage. Further, damage-induced increases in Brca2 mobility are accompanied by an increase in binding to its cargo, Rad51, in the nucleoplasm. Thus, release from functional sequestration into a freely diffusing nucleoplasmic pool may increase the availability of Brca2 for its interaction with Rad51. This change could involve Brca2 release from an intermolecular interaction that prevents Rad51 binding and/or an increased chance of encounter with Rad51 molecules in the nucleoplasm.
Our findings suggest that Brca2 diffusion, a basic physicochemical property of the protein, is dynamically regulated by binding to partner molecules in the cell nucleus. Biochemical fractionation of cell lysates shows that a proportion of detergent-extractable Brca2 exists in stable macromolecular complexes of >1 MDa (17). Whether these complexes represent biologically relevant states that exist in living cells is uncertain, but their existence is consistent with models for Brca2 diffusion that include a slow-diffusing component. The sheer size of these Brca2-containing complexes, and/or the presence of elements that transiently tether them to structures like the nuclear matrix, could help explain their restricted mobility. In this light, our finding that DNA damage induces the redistribution of Brca2 into simpler forms with higher mobility could reflect release from this type of functional sequestration. The identification of binding partners that mediate sequestration and release will be important in understanding the regulation of this critical tumor suppressor pathway. Potential candidates include the FANC complex, PALB2, EMSY, and DSS1 (4), known binding partners of Brca2 that participate in HR.
Regulated protein redistribution across microscopically visible nuclear compartments has previously been described (29). The noninvasive nature of FCS and its selectivity for the soluble fraction of a protein have permitted us to reveal a mechanism of sequestration of a nuclear protein into slow-moving complexes within the diffusing pool (rather than, for example, into static structures visible under conventional light microscopy), from which it can be released, in a regulated fashion, during a biological process. Such a mechanism may be particularly relevant in the case of nuclear proteins that assemble at specific sites on DNA, when diffusion limits the efficiency of DNA scanning (30, 31). Thus, the increase in Brca2 mobility may augment the probability of encounters with immobile DNA breaks (2) within the nuclear volume. It is tempting in this light to speculate that signal-initiated changes in the diffusion of nucleoplasmic proteins may provide a general means to control other essential DNA transactions, including replication, repair, or transcription.


FCS Calibration and Acquisition.

Fluorescence correlation spectroscopy measurements were performed using the confocor2 system as described previously (8). The pinhole of the FCS detector was aligned using 10 nM Rhodamine 6G (Rh6G) solution excited at 488 nm and imaged over the 505- to 550-nm spectral band. The same setup was used to measure fluctuations in the fluorescence emission of EGFP. Using a 70-μm pinhole for the FCS detector, the optimal excitation intensity for Brca2.EGFP cells was estimated at 75 μW by preliminary experiments. Ten sequential readings of 5–10 s each were performed per cell; the first sequence was discarded to exclude artifacts generated by immediate photobleaching of immobile molecules. Fluctuation traces exhibiting gross deviations in the count rate (suggestive of cell movement) or a poor signal-to-noise ratio (less than three times the background count rate from DT40 nuclei not expressing fluorophore-tagged proteins) were discarded. For FCS analysis of TMR-dextran and Cy3-Oligo(dT), the system was calibrated using a solution of free tetramethyl rhodamine (TMR) (10 nM), for an optical setup of 543 nm excitation and 560 LP emission.

Cross-Correlation Acquisition.

Fluorescence cross-correlation measurements were performed using the Zeiss Confocor2 system, using an optical arrangement for study of EGFP and mCherry—i.e., excitation by 488- and 543-nm lasers, appropriate beamsplitters, and detection bandpass filters 505–530 for EGFP [channel (Ch)2] and 600–650 for mCherry (Ch1). System setup and calibration were performed using a modification of the protocols described in ref. 7. Briefly, 10 nM Rh6G solution was used to calibrate the optimal position of the collimator and pinholes for maximum (max) count rate in both EGFP and mCherry channels. Then, the pinhole for the red channel (Ch1) was adjusted (diameter and x, y, z position), using excitation of Rh6G by the 488-nm laser, to optimize for maximum overlap. Cellular readings were performed on Brca2.EGFP Cherry.Rad51 cells, similar to FCS acquisition. A single LSM image (max speed, no averaging) was acquired to check for cells that expressed sufficient quantity of the green and red fluorophore. The crosshair for FCCS measurement was positioned on the basis of the brightfield image, at the center of the nucleus. Ten 5-s readings were recorded from each cell.

Data Analysis.

The fluorescence fluctuations (δF) are converted within the confocor software into the autocorrelation function (G(τ)) by the equation
and into the cross-correlation function (Ggr(τ)) by the equation
where g and r represent the green and red channels, respectively. The average curves from multiple cells were compiled within the confocor software and exported to GraphPad Prism 5.0 for graphical depiction. Models and analysis regimens used for fitting the autocorrelation and cross-correlation functions are discussed in detail in SI Methods.


We thank Drs. S. Takeda (Kyoto University, Kyoto) and K. J. Patel (Medical Research Council Laboratory of Molecular Biology, Cambridge, UK) for providing DT40 reagents and protocols, Drs. K. Sato and M. Lee for technical assistance, Dr. Enrico Gratton (University of California, Irvine, CA) for critical input on the data analysis, and Drs. V. Wickramasinghe and M. Garnett for thoughtful comments. A.D.J. thanks Klaus Weisshart (Carl Zeiss GmBH, Jena, Germany) and the instructors of the European Molecular Biology Organization course (advances in high-resolution microscopy; Buenos Aires, 2006) for assistance with FCS methods. A.D.J. received a Gates Cambridge Scholarship and a Career Development Fellowship from the United Kingdom Medical Research Council, E.R. and R.M. received studentships from the Medical Research Council and the Wellcome Trust, respectively, and A.E. received funding from the Engineering and Physical Sciences Research Council (EP/F044011/1). The Medical Research Council supports work in A.R.V.’s laboratory.

Supporting Information

Supporting Information (PDF)
Supporting Information


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Published in

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Proceedings of the National Academy of Sciences
Vol. 107 | No. 50
December 14, 2010
PubMed: 21098284


Submission history

Published online: November 22, 2010
Published in issue: December 14, 2010


  1. DNA damage response
  2. protein dynamics
  3. fluorescence spectroscopy
  4. single-molecule imaging


We thank Drs. S. Takeda (Kyoto University, Kyoto) and K. J. Patel (Medical Research Council Laboratory of Molecular Biology, Cambridge, UK) for providing DT40 reagents and protocols, Drs. K. Sato and M. Lee for technical assistance, Dr. Enrico Gratton (University of California, Irvine, CA) for critical input on the data analysis, and Drs. V. Wickramasinghe and M. Garnett for thoughtful comments. A.D.J. thanks Klaus Weisshart (Carl Zeiss GmBH, Jena, Germany) and the instructors of the European Molecular Biology Organization course (advances in high-resolution microscopy; Buenos Aires, 2006) for assistance with FCS methods. A.D.J. received a Gates Cambridge Scholarship and a Career Development Fellowship from the United Kingdom Medical Research Council, E.R. and R.M. received studentships from the Medical Research Council and the Wellcome Trust, respectively, and A.E. received funding from the Engineering and Physical Sciences Research Council (EP/F044011/1). The Medical Research Council supports work in A.R.V.’s laboratory.


This article is a PNAS Direct Submission.



Anand D. Jeyasekharan
The Medical Research Council Cancer Cell Unit, Hutchison/Medical Research Council Research Centre, Cambridge CB2 0XZ, United Kingdom; and
Nabieh Ayoub2
The Medical Research Council Cancer Cell Unit, Hutchison/Medical Research Council Research Centre, Cambridge CB2 0XZ, United Kingdom; and
Present address: Department of Biology, Technion-Israel Institute of Technology, Haifa, Israel, 32000.
Robert Mahen2
The Medical Research Council Cancer Cell Unit, Hutchison/Medical Research Council Research Centre, Cambridge CB2 0XZ, United Kingdom; and
Jonas Ries
Eidgenössiche Technische Hochschule Zurich, Laboratory of Physical Chemistry, CH-8093 Zurich, Switzerland
Alessandro Esposito
The Medical Research Council Cancer Cell Unit, Hutchison/Medical Research Council Research Centre, Cambridge CB2 0XZ, United Kingdom; and
Eeson Rajendra
The Medical Research Council Cancer Cell Unit, Hutchison/Medical Research Council Research Centre, Cambridge CB2 0XZ, United Kingdom; and
Hiroyoshi Hattori
The Medical Research Council Cancer Cell Unit, Hutchison/Medical Research Council Research Centre, Cambridge CB2 0XZ, United Kingdom; and
Rajan P. Kulkarni
The Medical Research Council Cancer Cell Unit, Hutchison/Medical Research Council Research Centre, Cambridge CB2 0XZ, United Kingdom; and
Present address: David Geffen School of Medicine, University of California, Los Angeles, CA 90095.
Ashok R. Venkitaraman4 [email protected]
The Medical Research Council Cancer Cell Unit, Hutchison/Medical Research Council Research Centre, Cambridge CB2 0XZ, United Kingdom; and


To whom correspondence should be addressed. E-mail: [email protected].
Author contributions: A.D.J. and A.R.V. designed research; A.D.J., N.A., R.M., E.R., R.P.K., and H.H. performed research; N.A. and A.D.J. contributed new reagents/analytic tools; A.D.J., J.R., A.E., and A.R.V. analyzed data; and A.D.J., A.E., and A.R.V. wrote the paper.
N.A. and R.M. contributed equally to this work.

Competing Interests

The authors declare no conflict of interest.

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    DNA damage regulates the mobility of Brca2 within the nucleoplasm of living cells
    Proceedings of the National Academy of Sciences
    • Vol. 107
    • No. 50
    • pp. 21231-21943







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