SUPPLEMENTARY INFORMATION

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1 SUPPLEMENTARY INFORMATION DOI: /NNANO Role of cell cycle on the cellular uptake and dilution of nanoparticles in a cell population Jong Ah Kim, Christoffer Åberg, Anna Salvati, Kenneth A. Dawson Figure S1 Cell population context and its effect on nanoparticle uptake. a, A549 cell culture growth curves (two independent experiments) constructed as described in Methods. Regions 1, 2 and 3 indicate three recognisable stages of cell culture growth: exponential growth, confluence and cell death, respectively. By fitting the exponential part of the curve (solid line), a cell population doubling time of 20 hours can be derived, in good agreement with the 22 hours given by the supplier (ATCC). Unless otherwise stated, all experiments in this study were performed with cultures in the exponential growth phase, (seeding density 8,800 cells/cm 2, as indicated by the horizontal dashed line). Error bars represent standard deviation of three independent replicates. b, Nanoparticle uptake (40 nm yellow-green PS-COOH 25 µg/ml in cmem) by A549 cells seeded at different cell densities, as indicated in the figure. The data are the mean cell fluorescence intensities as a function of time obtained by flow cytometry. Error bars represent standard deviation of three independent replicates. The results indicate a lower nanoparticle uptake by cells seeded at a high cell density (53,000 cells/cm 2 ), possibly due to cell confluence effects. NATURE NANOTECHNOLOGY 1

2 Figure S2 Effect of PS-COOH nanoparticles on cellular metabolism. a, Intracellular ATP content of nanoparticle-treated A549 cells after 24 hours of exposure to different doses of 40 nm yellow-green PS-COOH nanoparticles in cmem, compared to the ATP content of untreated cells. The ATP content was measured as described in the Methods and the values are normalised with respect to the ATP content of untreated cells, averaged over three replicates with the error bars representing the standard deviation. The unchanged intracellular ATP levels of particle-treated cells compared to untreated cells, suggest that the nanoparticles do not impair the energetic metabolism of the cells. b, Cell count of nanoparticle-treated and untreated A549 cells as a function of time. A549 cells were exposed to 25 µg/ml 40 nm yellow-green PS-COOH in cmem for up to 72 hours. Cell numbers were counted at different exposure times (5, 24, 48 and 72 hours of exposure), as described in the Detailed Methods. The similar number of cells between particle-treated and untreated cells confirms that the nanoparticles do not impair cell viability. 2

3 Figure S3 Effect of nanoparticle uptake on cell cycle. In order to analyse the effect of nanoparticle uptake on the cell cycle, DNA staining with propidium iodide was performed, as described in the Methods, on untreated A549 cells and compared to that of cells incubated with 25 and 100 µg/ml 40 nm PS-COOH, 25 µg/ml 100 nm PS-COOH and 100 µg/ml 50 nm silica nanoparticles for the indicated times. The cell cycle distribution was not affected by nanoparticle uptake for the different concentrations, nanoparticle sizes and materials tested. 3

4 Figure S4 Concentration dependence of nanoparticle uptake. A549 cells were exposed to 40 nm yellow-green PS-COOH at 25 and 100 µg/ml in cmem for up to 28 hours, prior to cell fluorescence measurement by flow cytometry. The results show the mean cell fluorescence intensity obtained by flow cytometry as a function of exposure time. The uptake in cells exposed to a higher extracellular concentration of nanoparticles (100 µg/ml) is higher than in cells exposed to a lower concentration (25 µg/ml). These results suggest that the levelling off seen in Fig. 1b (for cells exposed to 25 µg/m nanoparticles) is not due to the cells being saturated with nanoparticles. 4

5 Figure S5 Definition of cells in the different cell cycle phases according to EdU labelling and 7-AAD staining. The total DNA of A549 cells was stained with 7-AAD, and the cells exposed to EdU for 30 min, as described in the Methods, prior to cell fluorescence measurement by flow cytometry. EdU is a nucleoside analogue: when cells are exposed to EdU, the cells performing DNA synthesis incorporate it in the newly synthesised DNA. The results show the two-dimensional distribution of cells in terms of their total DNA content (7- AAD) and the content of synthesised DNA (EdU) as a colour-map. Cells in S phase synthesise DNA and therefore exhibit a high EdU fluorescence intensity. G0/G1 and G2/M do not synthesise DNA, and therefore exhibit a low EdU fluorescence intensity, but can be resolved based upon their total DNA content (single and double, respectively). These considerations lead to a definition of cells in the different phases as shown by the indicated regions. On this basis, the fraction of cells in the different phases can also be calculated. 5

6 Figure S6 Convergence of the mean cell fluorescence intensities of cells in the different cell cycle phases, as a function of the number of analysed cells. A549 cells were exposed to 25 µg/ml 40 nm yellow-green PS-COOH in cmem, prior to cell fluorescence measurement by flow cytometry as described in the Methods. The data corresponding to the 28 hour timepoint shown in Fig. 1c were analysed by selecting only the first number of cells of the full data set and resolving the contribution of the different cell cycle phases in this subpopulation for increasing size of the subpopulation (ultimately all the analysed cells). a, Convergence of the mean fluorescence intensities of the cells in the different cell cycle phases, as well as the mean fluorescence of all the cells. The mean values have been normalised by their respective final values. This analysis shows that the presented means (over all cells and over cells in the different cell cycle phases) are significant, because the values do not depend on the number of analysed cells. b, Convergence of the ranking of the mean fluorescence intensities of cells in the different cell cycle phases. In this case, the mean values were normalised by the mean fluorescence of all the cells. This analysis shows that the observed ranking in fluorescence of cells in the different cell cycle phases is significant. Indeed, already after acquiring the cell fluorescence intensity of less than 1,000 cells, the mean fluorescence intensity of cells in the different cell cycle phases is ranked in increasing order G2/M > S > G0/G1. 6

7 Figure S7 Ranking of the fluorescence intensity of cells in the different cell cycle phases in different cell lines. Different cell types were exposed to 40 nm yellow-green PS-COOH (25 µg/ml in complete DMEM for SH-SY5Y neuroblastoma cells and 1321N1 astrocytoma cells or complete EBM-2 medium in the case of hcmec/d3 cells) for up to 28 hours. After exposure to nanoparticles, the cells were stained with EdU and 7-AAD in order to determine the different cell cycle phases and measure their fluorescence intensity by flow cytometry as described in the Methods. The plots show the mean cell fluorescence intensities of all cells and the cells in G0/G1, S and G2/M phases of the cell cycle, defined by EdU/7-AAD doublestaining as shown in Supplementary Fig. S5. a, SH-SY5Y neuroblastoma cells (doubling time 48 hours). b, 1321N1 astrocytoma cells (doubling time 22 hours). c, hcmec/d3 endothelial cells (doubling time estimated to be 27 hours). For all the investigated cell types, thus also for different doubling times and cell culture media, the mean fluorescence intensity of cells in the different phases was ranked in the order G2/M > S > G0/G1. 7

8 Figure S8 Ranking of the fluorescence intensity of cells in the different cell cycle phases after treatment with different nanoparticles. a-d, A549 cells were treated for 24 hours with, respectively, 25 µg/ml 40 nm yellow-green PS-COOH in serum-free medium, 100 µg/ml 40 nm yellow-green PS-COOH in complete medium, 25 µg/ml 100 nm yellow-green PS-COOH in complete medium or 100 µg/ml 50 nm green silica in complete medium. After exposure to nanoparticles, the cells were stained by propidium iodide in order to determine the different cell cycle phases and their fluorescence intensity measured by flow cytometry as described in the Methods. It should be noted that in the case of panel a, where cells were incubated in serum free conditions for 24 hours, synchronisation in G0/G1 phase was observed. This synchronisation is also observed for untreated cells under the same serum-free conditions (as shown in Fig. 4a). The fluorescence intensity of the cells in the different cell cycle phases ranked in the order G2/M>S>G0/G1 regardless of nanoparticle concentration, size and material (thus with different protein corona) and the same ranking was confirmed also in absence of proteins (serum-free conditions). 8

9 Figure S9 Cell fluorescence intensity distributions simulated assuming asymmetrical partitioning of the intracellular load upon cell division. In the simulations the two daughter cells were assumed to split the intracellular nanoparticle load such that one daughter took 30% and the other 70% of the nanoparticles internalised by the parent cell. a-c, Distributions corresponding to those in Fig. 1e, the only difference being the asymmetrical partitioning. The mean accumulation of cells in the different cell cycle phases is not affected by the asymmetrical partitioning, and hence the corresponding averages are the same as those shown in Fig. 1f. The results show that eventual asymmetry would not be possible to discern in the broad fluorescence distributions. 9

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11 Figure S10 Dilution of intracellular nanoparticle load due to cell division (full time course of the experiment reported in Fig. 2). A549 cells were exposed to 40 nm yellow-green PS- COOH (25 µg/ml in cmem) for 4 hours and subsequently washed with PBS to remove the nanoparticle-containing media. The S phase cells were then labelled with EdU, as described in the Methods, and the cells grown further. At different times after the initial labelling, EdU and 7-AAD staining was performed as described in the Methods, followed by cell fluorescence measurement by flow cytometry. a, Mean nanoparticle fluorescence intensity of all the cells, obtained by flow cytometry, as a function of time after EdU-labelling. Note the log-scale. The solid red line shows a fit to the data corresponding to a splitting ratio of b, Two-dimensional distribution of cells in terms of their total DNA content (7-AAD) and the content of synthesised DNA (EdU) as a colour-map. EdU-labelled cells (originally in S phase at the time of labelling) with double DNA content are close to dividing ( Before division ). After division these cells will have single DNA content and reduced EdU content ( After division ). In this way, the evolution of the EdU-labelled cells can be followed. c-e, Corresponding nanoparticle fluorescence, EdU and total DNA content distributions of the EdU-labelled cells before (red) and after (blue) division. These results show the simultaneous splitting of nanoparticle load, EdU and DNA content upon cell division. 11

12 Figure S11 Effect of EdU incorporation on the total DNA content distribution. In order to ensure that the incorporation of EdU into newly synthesised DNA does not affect cell cycle progression, S phase A549 cells were labelled with EdU as described in the Methods, and subsequently grown further. At different times after the initial labelling, EdU and 7-AAD staining was performed as described in the Methods, followed by cell fluorescence measurement by flow cytometry. A549 cells that were not exposed to EdU were used as a control. The similarity between the DNA content distributions of EdU-labelled (right) and untreated control cells (left) suggests that EdU does not affect cell cycle progression (at least within the first 10 hours following EdU labelling). 12

13 Figure S12 Evolution of EdU-labelled cells through the cell cycle. S phase A549 cells were labelled with EdU as described in the Methods, and subsequently grown further in order to follow the evolution of EdU-labelled cells as they progressed through the cell cycle. At different times after the initial labelling, EdU and 7-AAD staining was performed as described in the Methods, followed by cell fluorescence measurement by flow cytometry. The two-dimensional distributions of cells in terms of their total DNA content (7-AAD) and the content of synthesised DNA (EdU) as a colour-map allow the determination of appropriate times when many of the EdU-labelled cells are in the S, G2/M and G0/G1 phases, respectively (see Supplementary Fig. S13 for the definition of the different phases). 13

14 Figure S13 Definition of cell cycle phases used to analyse the uptake rate during the different cell cycle phases (Fig. 3 and Supplementary Fig. S14). S phase A549 cells were labelled with EdU as described in the Methods, and then subsequently grown further in order to follow the evolution of EdU-labelled cells as they progressed through the cell cycle. At different times after the initial labelling, EdU and 7-AAD staining was performed as described in the Methods, followed by cell fluorescence measurement by flow cytometry. The results are presented as the two-dimensional distributions of cells in terms of their total DNA content (7-AAD) and the content of synthesised DNA (EdU) as a colour-map. The panels from left to right correspond to the three independent uptake studies performed 2, 6 and 12 hours after the initial EdU-labelling. The different cell cycle phases of the EdUlabelled cells were defined as indicated ( S cells, G2/M cells and G0/G1 cells), and the nanoparticle fluorescence of these cells analysed. The results are shown in Fig. 3. As an independent verification, the uptake during the different cell cycle phases was also compared to the uptake of the EdU negative cells (also indicated in this figure), and the results are shown in Supplementary Fig. S14. 14

15 Figure S14 Comparison of the nanoparticle uptake by cells in the different cell cycle phases with EdU-negative cells. S phase A549 cells were labelled with EdU as described in the Methods, and then subsequently grown further in order to follow their evolution as they progressed through the cell cycle. 2, 6 and 12 hours after the initial EdU-labelling, cells were exposed to 25 µg/ml 40 nm yellow-green PS-COOH in cmem for different times, prior to EdU and 7-AAD staining as described in the Methods. The mean cell fluorescence was then measured by flow cytometry, resulting in three independent uptake experiments. The results compare the uptake by cells in a given phase (as defined in Fig. S13) with the EdU-negative cells (also defined in Fig. S13). Error bars represent the standard deviation of three independent replicates. The results show that the uptake by cells in the different phases is comparable to the uptake by EdU-negative cells, consistent with the results shown in Fig

16 Figure S15 - Control of the bleed-through of EdU fluorescence into the nanoparticle channel. S phase A549 cells were labelled with EdU as described in the Methods, and then grown further for 6 hours in order to allow many EdU-labelled cells to enter the G2/M phase (as shown in Supplementary Fig. S12). Untreated cells and cells exposed to 25 µg/ml 40 nm yellow-green PS-COOH in cmem for different times were then stained for EdU and 7-AAD as described in the Methods, followed by cell fluorescence measurement by flow cytometry. The results are presented as the mean of the cell fluorescence intensity of the nanoparticle channel (green) obtained by flow cytometry, with error bars representing the standard deviation of three replicates. The fluorescence intensity in the nanoparticle channel (green) of the untreated EdU-labelled cells was very low in comparison to the values obtained for EdUlabelled cells exposed to the yellow-green PS-COOH nanoparticles. Moreover, the values remained constant for the duration of the experiment. The results exclude that the slightly higher green fluorescence of the EdU-labelled cells in G2/M phase shown in Fig. 3 (and Supplementary Fig. S14b) is due to increased bleed-through of the EdU fluorescence into the nanoparticle channel (green). 16

17 Figure S16 Forward/side scattering distribution of EdU-labelled G2/M cells and EdUnegative cells. S phase A549 cells were labelled with EdU as described in the Methods, and then grown further for 6 hours in order to allow many EdU-labelled cells to enter the G2/M phase (as shown in Supplementary Fig. S12). Then cells were exposed to 25 µg/ml 40 nm yellow-green PS-COOH in cmem for different times, and finally stained for EdU and 7- AAD as described in the Methods, followed by cell fluorescence measurement by flow cytometry. The top panels show the two-dimensional distributions of cells in terms of their total DNA content (7-AAD) and the content of synthesised DNA (EdU). The G2/M cells (red) and EdU-negative cells (blue) were defined as indicated (same definition as shown in Supplementary Fig. S13). The bottom panels show the two-dimensional distributions of cells in terms of their forward and side scattering, with the G2/M phase (red) and EdU-negative cells (blue) indicated. The results show that the forward scattering (an indicator of the size) of the cells in G2/M (red) is not different from that of the EdU-negative cells (blue). This suggests no major difference in size between cells in G2/M compared to EdU-negative cells, though, once in suspension, the forward scattering of the cells might not be sensitive enough for this purpose. Thus, the divergence between the uptake kinetics of cells in the G2/M phase and the EdU-negative cells (Supplementary Fig. S14) cannot be explained by a higher amount of residual nanoparticles adsorbed to the extracellular side of the plasma membrane, due to a larger surface area. A slightly higher side scattering is observed for the G2/M phase cells. 17

18 Figure S17 Nanoparticle accumulation curves simulated using various uptake rates for cells in the different phases of the cell cycle. a, Accumulation curves simulated using identical rates for all the cells (the same results as shown in Fig. 1f). b, Accumulation curves simulated using the measured rates for cells in the different cell cycle phases (the rates were obtained from the fits in Fig. 3). Note in particular that the slightly higher uptake rate during the G2/M phase shows only a minor quantitative effect due to G2/M being short in comparison with the other phases (as indicated by the low fraction of cells in G2/M in Supplementary Fig. S5). c, Accumulation curves simulated using a G2/M uptake rate 4 times larger than the measured one (Fig. 3). With this exaggerated rate there are significant effects, mainly when it comes to the accumulation in the G0/G1 and S phases, respectively; effects which are absent in the experimental data (shown in Fig. 1c). 18

19 Detailed Methods For uptake experiments, Click-iT EdU Flow Cytometry Kit (Invitrogen Corporation/Life Technologies Life Sciences, Carlsbad, CA) was used after exposure to nanoparticles to resolve the sub-population of cells in the different phases of the cell cycle. 150,000 cells were seeded on 35 mm diameter plates and were grown for 24 hours to allow cell adherence to the culture dish. Nanoparticle dispersions were prepared freshly under sterile conditions by diluting the stock in cmem (or the corresponding growth medium, depending on the cell line used in the experiment) to the required concentration, immediately prior to their addition to plates. Cells were incubated in approximately 1 ml of complete medium with nanoparticles at different concentrations (25 or 100 µg/ml) for different periods of time (30 minutes-72 hours). For the lowest concentration (25 µg/ml) of 40 nm PS-COOH the number of nanoparticles per cell was initially under these conditions, approximately halving during the first day, and reaching after 72 h. The saturated signal likely corresponds to roughly 10,000-50,000 internalised nanoparticles per cell, so that the total number of nanoparticles is in excess throughout the experiment. This conclusion was also corroborated by measuring the fluorescence (due to the nanoparticles) of the extracellular medium in the presence and absence of cells. Even though these measurements are affected by changes in ph, nanoparticle adhesion to the cell plate and media evaporation, only a 20% reduction could be measured after 72h, thus confirming that the experiments were performed in a regime of excess nanoparticles. Prior to fixation, plates were washed three times with PBS and then incubated with 10 µm EdU nucleoside analogue in complete media for 30 minutes at 37 C to label S phase cells. After EdU-labelling, cells were washed three times with PBS, harvested with 0.05% Trypsin-EDTA 1x, centrifuged for 3 minutes at 1500 rpm and 15 C, washed with 1% BSA in PBS, fixed, permeabilised (with Triton-X 100-based reagent) and treated for EdU detection according to the manufacturer s instructions. For total DNA staining, the Red Cell Cycle dye provided in the kit was used. Cells were finally resuspended in 0.5 ml of 1% BSA in PBS for their analysis on Dako CyAn-ADP flow cytometer equipped with 405 and 488 nm lasers. A total of 15,000 events were acquired per sample. Data were analysed using the Summit software (DAKO) and are presented as the mean cell fluorescence intensity of the so obtained distribution, averaged among the replicates, as specified in figure legends. Error bars represent the standard deviation among the replicates. To study the decay of an initial intracellular nanoparticle load, cells were incubated in approximately 1 ml of cmem with 25 µg/ml nanoparticles for 4 hours. After three PBS washes, Click-iT EdU Flow Cytometry Kit was used to label the sub-population of cells in S phase. Cells were pulse-labelled with 10 µm EdU nucleoside analogue in cmem for 30 minutes, then washed three times with PBS and further incubated in approximately 2 ml of EdU-free and nanoparticle-free cmem for various intervals of time. Cells were finally harvested, fixed and treated for EdU detection as explained above, also using the Red Cell Cycle dye for total DNA staining, prior to their analysis by flow cytometry. A total of 50,000 events were acquired per sample. For the analysis of uptake rates in the different cell cycle phases, Click-iT EdU Flow Cytometry Kit was used to label the sub-population of cells in S phase prior to exposure to nanoparticles. After pulse-labelling the cells with 10 µm EdU nucleoside analogue for 30 minutes and washing three times with PBS, cells were grown in cmem for 2, 6 and 12 hours 19

20 to allow the EdU-labelled subpopulation to enter respectively the S, G2/M and G1 phases. Cells were then treated with nanoparticles in EdU-free cmem for various time intervals as explained above. After exposure to nanoparticles, cells were harvested, fixed, treated for EdU detection and their total DNA stained with the Red Cell Cycle dye provided in the kit, before their analysis by flow cytometry. A total of 15,000 events were acquired per sample. To count the number of cells after exposure to nanoparticles, 35 mm plates were seeded with 150,000 A549 cells and after 24 hours, the cells were exposed to 25 µg/ml 40 nm yellow-green PS-COOH in cmem for up to 72 hours. Nanoparticle-treated samples were harvested at different exposure times (5, 24,48 and 72 hours of exposure), together with untreated cells prepared and grown for the same times in the same conditions (without exposure to nanoparticles). The cells were resuspended in 1ml of PBS and the cell number counted manually in a haemocytometer chamber. Trypan blue was used to stain dead cells and exclude them from the count. To study possible perturbations on the cell cycle, total DNA staining was performed. After incubation with nanoparticles (and/or EdU) cells were harvested as previously described and fixed with 70% ice-cold ethanol at 4 C for 1 hour. After centrifugation, cells were resuspended in 0.5 ml of PBS, treated with 50 µl RNAse 1 mg/ml for 5 minutes and stained with 25 µl propidium iodide (Sigma-Aldrich Fine Chemical Co., St. Louis, MO) 1 mg/ml. A total of 15,000 events were analysed by flow cytometry per sample. To analyse uptake at different degrees of confluence, 60 mm plates were seeded with different starting cell densities (40,000, 250,000 or 1,500,000 cells per plate) and after allowing 24 hours for adhesion, cells were incubated with 25 µg/ml nanoparticles in cmem for up to 24 hours. Cells were finally harvested, fixed with 70% ethanol and stained with propidium iodide as explained above for their analysis on flow cytometry. A total of 15,000 events were acquired per sample. 20

21 Supplementary Discussion Role of cell cycle on the cellular uptake and dilution of nanoparticles in a cell population Jong Ah Kim, Christoffer Åberg, Anna Salvati, Kenneth A. Dawson Numerical Simulations of Nanoparticle Uptake by A Cell Population Undergoing Exponential Growth At all times, cells in the G2/M phase have had the longest time to take up nanoparticles, followed by cells in the S phase; in contrast, cells that have just divided are in the G1 phase. This fact is certainly consistent with the experimentally observed ranking (Fig. 1c), but in order to assess if this effect is quantitatively large enough to explain the experimental observation, simulations were performed with an ensemble of cells under somewhat idealised circumstances. In the simulations, all cells were assumed to cycle at the same rate. Different cycle lengths can be included, but would likely not lead to quantitatively different results. The positions of individual cells along the cell cycle, their ages, were randomly chosen according to an exponentially decaying distribution, as illustrated in the figure below. This distribution is well-known in cell proliferation analysis 1,2,3, and leads to an exponentially growing population of cells. Exponentially decaying distribution along cell cycle with the individual phases indicated. Based on the distribution, the traversal time for a given phase can be related to the fraction of cells in that phase, as illustrated graphically. The shaded bins show a population of 10 6 cells randomly chosen according to the distribution and used as the initial population for the simulations. In the simulations, cells traversed the cell cycle with time. Since the exponentially decaying distribution (normalised for the total number of cells) is time-independent, the fraction of cells in each phase remains constant throughout the simulations. The time spent in each phase can then be related to the fraction of cells in a given phase 2,3, as illustrated in the figure above. The fact that the fraction of cells in each phase remained constant in the simulations implicitly assumes that the cell population is in a state of exponential growth and that effects due to confluence have not been taken into account. We can therefore only

22 compare the simulations with experiments for a limited time course, during which these assumptions remain approximately true. The simulations can be validated to data by using the nucleoside analogue EdU (5- ethynyl-2 -deoxyuridine) together with DNA staining (7-AAD) to follow experimentally the division of those cells that were in S phase at the time of labelling. Since the labelled (S phase) cells have to traverse G2/M prior to division, a comparison with the simulations can be performed by counting the number of cells that have divided in the simulations, after a delay corresponding to the time spent in G2/M. The prediction, using only parameters independently measured (fraction of cells in the different phases) or acquired (cell population doubling time), is compared with the experimental data in Fig. 2b to excellent agreement. As each simulated cell traversed the cell cycle, it was also assumed to take up nanoparticles with a given rate. In the simplest case (Fig. 1e-f), the rate was assumed independent of the cell s instantaneous position along the cell cycle. However, each cell was allowed to have its own rate, simulating a heterogeneity in the cell population when it comes to nanoparticle uptake. The rate with which an individual cell takes up nanoparticles was, furthermore, assumed to be inherited by both daughter cells upon cell division. A log-normal distribution of rates was used since it reproduces the appearance of the experimentally observed distributions measured by flow cytometry, though we imply no deeper significance than that. To investigate the effect of different rates of uptake during the different phases (Supplementary Fig. S17), simulations were performed where the uptake during a given phase was assumed constant, but different among the phases. Again each cell took up nanoparticles with its own rate; this rate being inherited by both daughter cells upon cell division. A 'fictitious rate',, was assigned to each cell from the same log-normal distribution as for the simpler case of constant rate during the cell cycle (previous paragraph). The rates of uptake during the different phases for a given cell,, and 2, respectively, were scaled from the 'fictitious rate' according to where denotes the measured rate and the fraction of cells in phase, respectively. The respective numerators in the expressions above imply that the ratios of the rates of uptake during the different phases of any given cell exactly match the ratios of the measured rates. In other words, a cell that takes up nanoparticles faster than average during the G0/G1 phase, also takes up nanoparticles faster than average during the S and G2/M phases. Furthermore, the rate of uptake for any given cell during G2/M phase is somewhat larger than during the other phases. The denominator in the above expressions was used for simplicity of comparison, since this normalisation ensures that the mean of the full population exactly matches that for the simpler case of constant rate of uptake during the different phases (previous paragraph). No export of nanoparticles was taken into account, and therefore the only diluting effect on the average number of internalised nanoparticles is cell division. Upon cell division, each simulated cell was in the simplest case (Fig. 1e-f and Supplementary Fig. S17) assumed to give exactly half of its intracellular load of nanoparticles to each daughter cell. To investigate the effect of asymmetrical partitioning in qualitative terms (Supplementary Fig. S9),

23 simulations were also performed where one of the daughter cells received 70% and the other 30% of the intracellular load of the parent cell. In the numerical evaluation, we used a cell cycle length of 22 hours (the cell population doubling time given by the supplier), and fractions of cells in the different phases of 52% G0/G1, 40% S and 8% G2/M, respectively, as determined experimentally from the EdU and 7-ADD double staining (Supplementary Fig. S5). The initial population consisted of 10 6 cells (growing with time), but results were re-normalised to correspond to the 15,000 cells measured experimentally. The means corresponding to the different phases were calculated by averaging over the cells in a given phase, as defined by their position along the cell cycle (illustrated in the figure above). Results for the simplest case of equal rates of uptake during all phases and symmetrical split upon cell division are reported in Fig. 1: Figure 1f shows the average accumulations corresponding to the experiment shown in Fig. 1c, and Fig. 1e shows distributions of cell fluorescence intensity corresponding to the experiment shown in Fig. 1d. Supplementary Fig. S9 shows the effect of asymmetrical partitioning, keeping the rates equal during the different phases, on the cell fluorescence intensity distributions. Note that the average nanoparticle accumulation by cells instantaneously in a given phase is not affected by an asymmetrical partitioning, since the average partitioning is still half (for example, if one daughter receives 70% of the intracellular load and the other 30%, the average is still 50%). The averages corresponding to Fig. S9 are therefore the same as shown in Fig. 1f. In particular, this implies that the ranking of the phases is completely independent of eventual asymmetry of the partitioning. Supplementary Fig. S17 compares the average uptake with different rates of uptake in the different phases. Notably, the somewhat larger uptake rate measured during the G2/M phase (Fig. 3) does not affect the nanoparticle accumulation significantly, simply due to the fact that the time spent during G2/M is comparatively short. The observed ranking in nanoparticle accumulation for cells in the different phases is therefore most likely not due to the measured (minor) differences in uptake rates. In the absence of a nanoparticle source, the simulation predicts an exponential decay of the average number of nanoparticles per cell with a decay constant given by where is the cell population doubling time. Figure 2c shows the decay of the average fluorescence after loading cells with nanoparticles for some time, and subsequent removal of the nanoparticle source. A one-parameter fit (with initial fluorescence as a free parameter, and the decay constant fixed) shows that the simulation gives a good description of the experimental data (solid line in Fig. 2c). References 1. Gray, J. W. Cell cycle analysis from computer synthesis of deoxyribonucleic acid histograms. J. Histochem. Cytochem. 22, (1974). 2. Steel, G. G. Growth Kinetics of Tumors: Cell Population Kinetics in Relation to the Growth and Treatment of Cancer (Oxford Univ. Press, Oxford, 1977). 3. Montalenti, F., Sena, G., Cappella, P. & Ubezio, P. Simulating cancer-cell kinetics after drug treatment: Application to cisplatin on ovarian carcinoma. Phys. Rev. E 57, (1998).