Supplementary Information
“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/cm2, 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.
1
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
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
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
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 (7AAD) 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
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
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
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
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.
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10
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 PSCOOH (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 0.62.
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.
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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
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
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
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. 3.
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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).
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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 7AAD 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.
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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).
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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 5∙106 under these conditions, approximately halving
during the first day, and reaching 5∙105 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
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.
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