Cellular Uptake and Intracellular Trafficking of Poly(N-(2

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Cellular Uptake and Intracellular Trafficking of Poly(N-(2-Hydroxypropyl) Methacrylamide) Claudia Battistella, Romain Guiet, Olivier Burri, Arne Seitz, Stephane Escrig, Graham W. Knott, Anders Meibom, and Harm-Anton Klok Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.8b01372 • Publication Date (Web): 05 Nov 2018 Downloaded from http://pubs.acs.org on November 8, 2018

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Biomacromolecules

Cellular Uptake and Intracellular Trafficking of Poly(N-(2-Hydroxypropyl) Methacrylamide)

Claudia Battistella,1 Romain Guiet,2 Olivier Burri,2 Arne Seitz,2 Stéphane Escrig,3 Graham W. Knott,4 Anders Meibom3,5 and Harm-Anton Klok1,*

1

École Polytechnique Fédérale de Lausanne (EPFL), Institut des Matériaux et

Institut des Sciences et Ingénierie Chimiques, Laboratoire des Polymères, Bâtiment MXD, Station 12, CH-1015 Lausanne, Switzerland. 2

École Polytechnique Fédérale de Lausanne (EPFL), Faculté des sciences de la

vie, Bioimaging and optics platform, Bâtiment AI, Station 15, CH-1015 Lausanne, Switzerland. 3Laboratory

for Biological Geochemistry, School of Architecture, Civil and

Environmental Engineering, Ecole Polytechnique Fédérale de Lausanne (EPFL), CH-1015 Lausanne, Switzerland. 1 ACS Paragon Plus Environment

Biomacromolecules 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

4

École Polytechnique Fédérale de Lausanne (EPFL), Faculté des sciences de la vie, Bioelectron Microscopy Core Facility, Bâtiment AI, Station 19, CH-1015 Lausanne, Switzerland.

5Center

for Advanced Surface Analysis, Institute of Earth Sciences, University of Lausanne, CH-1015 Lausanne, Switzerland.

CORRESPONDING AUTHOR: [email protected]; Tel: + 41 21 693 4866

2 ACS Paragon Plus Environment

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ABSTRACT

Cellular uptake and intracellular trafficking of polymer conjugates or polymer nanoparticles is typically monitored using fluorescence-based techniques such as confocal microscopy. While these methods have provided a wealth of insight into the internalization and trafficking of polymers and polymer nanoparticles, they require fluorescent labelling of the polymer or polymer nanoparticle. Since in biological media fluorescent dyes may degrade, be cleaved from the polymer or particle or even change uptake and trafficking pathways, there is an interest in fluorescent label-free methods to study the interactions between cells and polymer nanomedicines. This article presents a first proof-of-concept that demonstrates the feasibility of NanoSIMS to monitor the intracellular localization of polymer conjugates. For the experiments reported here, poly(N-(2-hydroxypropyl) methacrylamide)) (PHPMA) was selected as a prototypical polymer – drug conjugate. This PHPMA polymer contained a 19Flabel at the -terminus, which was introduced in order to allow NanoSIMS analysis. Prior to the NanoSIMS experiments, the uptake and intracellular trafficking of the polymer was established using confocal microscopy and flow cytometry. These experiments not only provided detailed insight into the kinetics of these processes, but were also important to select time points for the NanoSIMS analysis. For the NanoSIMS experiments, HeLa cells were investigated that had been exposed to the PHPMA polymer for a period 4 or 15 hours, which was known to lead to predominant lysosomal accumulation of the polymer. NanoSIMS analysis of resin-embedded and microtomed samples of the cells revealed a punctuated fluorine signal, which was found to co-localize with the sulphur signal that was attributed to the lysosomal compartments. The localization of the polymer in the endolysosomal compartments was confirmed by TEM analysis on the same cell samples. The results of this



study illustrate the potential of NanoSIMS to study the uptake and intracellular trafficking of polymer nanomedicines. INTRODUCTION The use of polymer conjugates and polymer nanocarriers for drug delivery can help to increase plasma half-life and drug bioavailability1,2,3 In case of cancer chemotherapy, the enhanced permeation and retention (EPR) effect provides a potential strategy towards passive tumor targeting. Translating the EPR effect into an effective clinical tool to treat human cancers, however, has proven a great challenge.4,5 Although enhanced drug bioavailability and efficient tumor targeting are important to maximize the efficacy of (anticancer) drugs and to reduce side effects, many drug targets are located in specific subcellular compartments. As a consequence, intracellular transport is another factor that contributes to the therapeutic outcome. After extravasation in the cancer tissue, polymers are typically internalized by endocytosis.6,7 This process starts with the engulfment of the cargo by the cell membrane and results in the formation of intracellular vesicles called early endosomes, which are characterized by a slightly acidic pH. The internalized material is then shuttled from the early endosomes to the slightly higher acidic late


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endosomes, before being finally sorted to the lysosomes, where enzymatic digestion takes place.8 By conjugating drugs via pH- or reduction-sensitive bonds or via enzymatically cleavable linkers to a polymer carrier or by using polymer nanoparticles that disintegrate upon change of pH, redox environment or upon exposure to intracellular enzymes, these physicochemical and biochemical stimuli can be used to promote drug release in specific subcellular locations.9 First developed by Kopeček and Duncan in the 1970s, poly(N-(2-hydroxypropyl) methacrylamide) (PHPMA)-based carriers have been extensively explored for the intracellular delivery of anticancer drugs such as doxorubicin, camptothecin, paclitaxel and platinates.10,11,12,13,14

The interest in exploring PHPMA-based

conjugates as chemotherapeutics has also triggered a need to study and understand the internalization and intracellular trafficking of these nanomedicines. Using fluorescent-labeled samples, the cellular internalization of PHPMA conjugates has been studied using fluorometry or flow cytometry.15-18 Alternatively, radioactive labels and radioassays have also been used.19 While these techniques allow to quantitatively assess cellular uptake, they do not provide insight into the intracellular trafficking pathways of the polymer conjugates. Intracellular trafficking of PHPMA 5 ACS Paragon Plus Environment


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conjugates has been studied using confocal microscopy.17-23 Initially, these approaches were based on the subcellular localization of fluorescent drugs. For instance, 6 hours exposure of cells to a PHPMA-mesochlorin e6 (Mce6) conjugate resulted in polymer accumulation in the lysosomes.20 Similarly, confocal microscopy was used to monitor the cleavage of a lysosomal sensitive linker (GFLG) and subsequent doxorubicin (Dox) release from PHPMA conjugates.19 Duncan and coworkers combined subcellular fractionation techniques with confocal microscopy to determine

Dox

release

from

a

PHPMA-GFLG-Dox

conjugate.21

In

these

experiments, 5 hours incubation of cells with the polymer conjugate was found to result in both punctuate cytoplasmic doxorubicin fluorescence as well as a disperse signal, which was attributed to doxorubicin that had been released from the carrier. Jensen et al. monitored time-dependent intracellular trafficking by using PHPMA-dye conjugates. This study, however, did not use endosomal and lysosomal fluorescent markers, which would have allowed co-localization studies with the endocytic vesicles.18 Other reports have studied the effects of antibody conjugation or insertion of charged and hydrophobic moieties on the cellular internalization and accumulation in the late endosomal vesicles of PHPMA copolymers.17,22 Although these studies 6 ACS Paragon Plus Environment

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have given valuable insight into the endocytic trafficking of PHPMA conjugates, the investigation of the complete time-frame of the endocytic process, from early endosomal uptake to lysosomal accumulation, would give precise information on the time-dependent

exposure

of

the

polymer

to

the

different

endolysosomal

environments. All of the examples discussed above have relied on the use of fluorescent-labeled polymers. While these experiments have provided significant insight, there are also a number of drawbacks and limitations related to the use of fluorescent labeled samples. First, exposure of the polymer-dye conjugates to the highly degradative cellular environment can result in the cleavage of the fluorescent dye from the polymer carrier.24-26 Second, the fluorescent label can interfere with biological processes27 and finally, concentration- or pH-dependent dye quenching might lead to additional artifacts.24 Additional challenges arise when this approach is used to monitor not only intracellular trafficking but also drug release. In this case, both polymer and drug need to be tracked at the subcellular level. To this end, unless an intrinsic fluorescent drug is used, the drug needs to be substituted with a fluorescent dye. Fluorescent carrier and drug need to be monitored simultaneously and co7 ACS Paragon Plus Environment


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localization studies with several stained organelles can result in complicated assays. Moreover, reported co-localization studies generally lack quantitative information.7,26 Experimental techniques that do not require fluorescent labels are attractive tools that can be used to complement, i.e. further refine or quantify, the insights obtained from confocal microscopy. One example of a powerful, fluorescent label-free technique that can be used to study cellular internalization and intracellular trafficking is Nanoscale Secondary Ion Mass Spectrometry (NanoSIMS). NanoSIMS is an attractive technique since it offers high spatial resolution (down to 50 nm) and the possibility to obtain quantitative information while also excluding the need for bulky labels.28 NanoSIMS allows the semi-quantitative analysis of trace elements and isotopes by sample erosion via a focused ion primary beam.28 NanoSIMS has been used to track organometallic anticancer drugs in cells.29-32 Proetto et al. combined fluorescence microscopy with NanoSIMS to image

15N-labeled

fluorescent

nanoparticles (d = 130 – 140 nm) that carried a platinum-based anticancer drug.33 In this case, fluorescence microscopy allowed subcellular carrier co-localization with endocytic vesicles, while NanoSIMS analysis was used to simultaneously track both internalized carrier and drug. 8 ACS Paragon Plus Environment

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While this technique, in combination with fluorescence microscopy, has been successfully used to detect polymer nanoparticles, NanoSIMS has not yet been explored to directly monitor the intracellular localization and distribution of polymers or polymer conjugates. The aim of this study is to assess the feasibility of NanoSIMS to monitor the cellular uptake and intracellular trafficking and localization of polymer conjugates, using PHPMA as a model system. In order to define the timeframe of these experiments, the NanoSIMS studies were preceded by detailed confocal microscopy experiments in order to map the kinetics of the cellular uptake and intracellular trafficking of PHPMA. As pointed out above, there have been prior confocal microscopy studies on PHPMA polymers, but the aim of most of these studies was to visualize cellular uptake of the polymer or final accumulation of the polymer in the lysosomes rather than the detailed kinetics of these processes. The time dependent cellular internalization experiments laid the ground for the NanoSIMS experiments by allowing to determine the time window (4 – 15 hours) that results in predominant lysosomal accumulation of the PHPMA polymers. HeLa cells that were exposed to the PHPMA polymer for 4 or 15 hours were then investigated 9 ACS Paragon Plus Environment


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with NanoSIMS to identify the subcellular location and distribution of the fluorescent label-free polymer and to assess the feasibility of this technique, alone or in combination with TEM, to track and localize such polymers and polymer conjugates at the subcellular level.

EXPERIMENTAL SECTION MATERIALS Annexin V-Alexa Fluor 647 conjugate, Annexin buffer, CellLight lysosomes-green fluorescent protein (GFP), CellLight early endosomes-GFP, Rhodamine Red C2 maleimide and CellTrace Violet were purchased from Life Technologies. 1 mM Staurosporine

solution

in

DMSO,

2-(4-amidinophenyl)-6-indolecarbamidine

dihydrochloride (DAPI), poly(D-lysine) hydrobromide, tris(2-carboxyethyl)phosphine hydrochloride (TCEP), 1-amino-2-propanol (HPA), sodium borohydride (NaBH4), Sephadex G 15 and 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU) were purchased from Sigma-Aldrich. triethylamine

Pentafluorophenol (Et3N)

Trifluoropropylamine

and

was

1,4-dioxane

hydrochloride

was

purchased were

from

from

obtained


MatrixScientific

Acros

from

Organics.

Apollo

and 3,3,3-

Scientific

and

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dimethylformamide (DMF) from VWR. DMF was dried using a Pure Solv solvent purification system. The -19F, -Rhodamine Red labeled PHPMA polymer investigated in this study was prepared as described previously.34 Figure S1 and Figure S2 present

19F-

and 1H-NMR spectra of the -19F-labeled PPFMA precursor.

Figure S3 shows the calibration curve that was used for the spectrophotometric determination of the degree of Rhodamine red end functionalization. The 1H-NMR spectrum of the -19F-labeled modified PHPMA is included in Figure S4. Table 1 summarizes the number-average molecular weights and dispersities (Mw/Mn) of the precursor -19F-labeled PPFMA precursor and the -19F, -Rhodamine Red labeled PHPMA.

METHODS NMR spectroscopy. 1H- and

19F-NMR

spectra were recorded on a Bruker AV III-400

instrument at room temperature. 1H-NMR spectra of polymers were recorded with a relaxation time (d1) of 10 seconds and at least 256 scans using CD3OD and CDCl3 as solvents. Chemical shifts are reported relative to the residual proton signal of the solvent. 11 ACS Paragon Plus Environment


Size exclusion chromatography (SEC). Size exclusion chromatography (SEC) of PPFMA in THF was performed using an Agilent 1260 infinity system equipped with a Varian 390-LC refractive index (RI) detector, two PLgel 5 m Mixed C (Agilent) columns and a PLgel guard column. THF was used as eluent with a flow rate of 1 mL/min and the temperature was 40 °C. Samples were analyzed using conventional calibration with polystyrene (PS) standards ranging from 4910 Da to 549 KDa.

UV-Vis spectroscopy. The incorporation of Rhodamine Red C2 maleimide in the polymer was quantified using a Varian Cary 100 Bio spectrometer using quartz cuvettes.

Flow cytometry. Flow cytometry was performed using a LSRII cytometer equipped with violet (405 nm), blue (488 nm), green (561 nm) and red (640 nm) lasers and the data were analyzed using FlowJo software.


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Confocal microscopy. Confocal microscopy images were acquired on a Zeiss LSM700 microscope with a Plan-Apochromat 63x/1.40 Oil objective and a zoom factor of 1.4 in order to obtain 70 nm pixel in the image. Images were acquired sequentially in order to avoid excitation and emission bleed-through with the following settings for the individual channels: Ch1 400 nm – 470 nm, Ch2 595 nm – 560 nm, Ch3 560 nm – 700 nm. The pinhole was set to 1 AU for Channel 3 corresponding to an optical thickness of 0.8 µm. For the other channels, the pinhole was set in order to obtain the same optical slice thickness as in Ch1. 4 to 6 Images were acquired per each incubation time and the experiments were replicated 3 times. Vesicles are a spherical 3D objects. Due to the optical sectioning functionality of the confocal microscope both polymer chains and vesicles resemble disks, which suggested to use an object based co-localization analysis. Unfortunately, due to the resolution limit of microscopy, individual objects could not always be obtained. Hence, the object based analysis relying on the measure of distances between closest neighbor objects would have been biased. For this reason, Mander’s coefficients were determined at the different time points.35 Cell-to-cell differences in GFP expression and different background levels exclude the usage of fixed cut-off values for all images. Histogram-derived methods were chosen based on visual inspection of the thresholded images. Moments36 was used for vesicular compartments and RenyiEntropy37 for polymer images respectively. The outline of the cell was drawn manually. This is necessary to account for variations in cell size and expression 13 ACS Paragon Plus Environment


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level of the fusion construct. To perform the analysis, the Fiji distribution of ImageJ,38 the JACoP plugin39 and a customized Action Bar “BIOP JACoP Tools”40 enabling batch processing, were used.

NanoSIMS. For the subcellular detection of the fluorine-labeled PHPMA, samples were analyzed using a NanoSIMS 50L instrument. For anion analysis, the NanoSIMS uses a Cs+ primary beam that can be scanned over a squared surface to produce elemental images. The probe size (full-width-half-maximum of the Cs+ beam) was ~ 150 nm and beam current on the sample surface was around 2 pA. Image size was 38 x 38 µm with a 256 x 256 pixel image resolution and a dwell time of 5000 s per pixel. The NanoSIMS detectors were set up to collect and

32S−

12C14N−, 19F−

secondary ions simultaneously at a mass resolution sufficient to avoid any

isobaric interference. Each image was acquired following the same protocol: a phase of pre-sputtering where a larger Cs+ beam was scanned over the surface in order to remove the gold layer from the area of interest and to implant Cs+ into the sample and an acquisition phase consisting of five successive scans over the surface during which data were collected and were later cumulated during the data reduction.


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Transmission electron microscopy. Resin-embedded cells cut at a thickness of 70 nm were imaged in a transmission electron microscope (Tecnai Spirit, FEI Company) at a beam voltage of 80 kV.

PROCEDURES Cell culture. HeLa cells were cultured in DMEM (high glucose) medium supplemented

with

10

%

fetal

bovine

serum

(FBS)

(Gibco)

and

1%

penicillin/streptomycin (Lifetech). Cells were maintained in a humidified atmosphere containing 5 % (v/v) of CO2 at 37 °C.

Cell viability assay and polymer internalization kinetics. Cell viabilities were assessed using Annexin V-Alexa Fluor 647/DAPI assay via flow cytometry. HeLa cells were seeded in 24-well plates at a density of 25000 cells per well and incubated in 450 L medium for 18 h. Untreated cells were used as negative control and cells treated with Staurosporine were used as positive control. To this end, 1 L of 1 mM Staurosporine stock solution was add to each mL of cells and cells were incubated for additional 24 h. Cells were incubated with 0.6 mg/mL 19F-PHPMA-Rhodamine for 24 h, 15 h, 4 h, 2 h, 40 min and 10 15 ACS Paragon Plus Environment


min. After that, cells were gently trypsinized, collected in 10 mL tubes and washed with PBS twice. Negative, untreated control cells were kept on ice until analysis. Positive control cells and cells treated with the polymer were re-suspended in 100 L Annexin buffer (1X) containing 1 g/mL DAPI at a density of 1 x 106 cells/mL. 5 L of Annexin-V-Alexa Fluor 647 was then added to each tube and cells were incubated at room temperature for 15 minutes. Untreated cells stained only with DAPI and only with Annexin V-Alexa Fluor 647 were also prepared and used as controls. After the incubation period, 400 L Annexin buffer was added to each tube and cells were kept in ice until analyzed by flow cytometry. Cells were analyzed using FL 9 channel for DAPI and FL 6 for Alexa Fluor 647. In order to determine polymer internalization kinetics, intracellular Rhodamine was detected using FL 2 channel. Two independent experiments were performed and 10.000 events were collected.

CellTrace Violet proliferation assay. Cells were grown and incubated with the polymers as described in the previous section. After trypsin treatment, cells were incubated at 37 °C for 20 min in 5 mL PBS containing 5 L CellTrace Violet stock solution in DMSO. After this time, 25 mL DMEM medium was added and the cells were incubated for further 5 minutes. Cells were centrifuged, resuspended in fresh medium and seeded. One sample of cells was centrifuged, resuspended in PBS and analyzed by flow cytometry using FL 9 channel for CellTrace Violet detection (Seeding time). After 18 hour, cells were treated with 0.6 mg/mL polymer solution for 24 h, 15 h, 4 h, 40 min and 10 min. For each incubation time, prior to adding the polymer solution, one sample of cells was trypsinized, washed with PBS and analyzed by flow cytometry. At the end of the time course experiment, cells incubated with the polymer solution for the different times were collected, washed with PBS and analyzed by flow cytometry (End time). Comparison between CellTrace Violet-cell fluorescence at the “Seeding time”, prior to each polymer addition, and at the end of the time-course experiment 16 ACS Paragon Plus Environment

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“End time” allowed the determination of the time-dependent cell division occurring during the experiment (see also Figure 3A).

Cell preparation for confocal microscopy. HeLa cells were seeded on 13 mm diameter sterilized poly(D-lysine)-coated coverslips in 24-well plates at a density of 25000 cells per well and incubated in 450 L medium for 18 h. After that, either 17 L (PPC 35) CellLight early endosomes GFP or 20 L (PPC 40) CellLight lysosomes GFP were added to the cell medium and the cells were further incubated for 24 h. During this period, the cell medium was replaced by free phenol red-DMEM medium containing 0.6 mg/mL

19F-PHPMA-Rhodamine

and the cells were incubated for 24,

15 h, 4 h, 2 h, 40 min and 10 min. For the 15 h incubation sample, the polymer solution was supplemented with the CellLight constructs in order to keep the cell exposure to the constructs constant among the different incubation times. At the end of the time-course experiment, cells were washed three times with DMEM without phenol red and twice with PBS and finally fixed using 2 % paraformaldehyde and 1 % glutaraldehyde solution in phosphate buffer (PB) (pH 7.4, 0.1 M) for one hour at room temperature. Cells were then washed twice with cacodylate buffer (pH 7.4, 0.1 M) and stained with 0.5 g/mL DAPI solution in the same buffer for 6 minutes. Cells 17 ACS Paragon Plus Environment


were washed four times and kept in cacodylate buffer during microscopy analysis. Three independent experiments were performed.

Sample preparation for TEM and NanoSIMS analysis. TEM and NanoSIMS analysis was performed on exactly the same samples that were studied with confocal microscopy. First, cells that had been investigated with confocal microscopy were post-fixed for 60 minutes in a solution of 1% osmium tetroxide and 1.5% potassium ferrocyanide in ice-cold 0.1 M cacodylate buffer. This was followed by staining with a solution of 0.1 g thiocarbohydrazide in 10 mL double distilled water for 20 min at room temperature. Subsequently, staining with 1% osmium tetroxide in 0.1 M cacodylate buffer solution was performed for 30 min. Then, the samples were dehydrated in an ascending alcohol series (1 X 50%, 1 X 70%, 2 X 96%, 2 X 100%, 3 minutes each) and embedded in Durcupan resin, which was then hardened overnight at 65 °C. 70 nm-thick sections were cut and placed on TEM finder grids. The grids and sections were gold-coated prior to NanoSIMS analysis to avoid charging effects.

RESULTS AND DISCUSSION 19F-PHPMA-Rhodamine

design and synthesis

To explore the feasibility of NanoSIMS to study cellular uptake and intracellular trafficking of PHMPA, the polymer needs to contain an isotope or trace element label. For this study,

19F

was selected as it is inexpensive as compared to isotopic

labelling, which is frequently used in NanoSIMS, and also is not present in biological 18 ACS Paragon Plus Environment

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environments, which helps to avoid interference with other intracellular compounds, Futhermore,

19F

gives rise to minimal steric alterations and possesses a high

metabolic stability.41 The

19F

end-labelled PHPMA was synthesized as outlined in

Scheme 1 following a previously published protocol.34 In brief,

19F-labelled

PHPMA

was obtained by RAFT polymerization of pentafluorophenyl methacrylate (PFMA) using a

19F-functionalized

poly(pentafluorophenyl

chain transfer agent to afford a -19F-functional

methacrylate)

(PPFMA)

polymer,

polymerization modification with hydroxypropylamine. The in this study did not only incorporate an

19F-label

followed

19F-labelled

by

post-

PHPMA used

at the -terminus of the polymer

chain, but was also functionalized at the -chain end with a Rhodamine Red fluorescent label, which was introduced by reaction of the thiol end group with a maleimide functionalized derivative of this dye. Table 1 summarizes the molecular weights and dispersities (Mw/Mn) of the

19F-PPFMA

and

determined by size exclusion chromatography and

19F-PHPMA

19F-NMR

polymers as

spectroscopy. The

degree of Rhodamine Red end-group functionalization of the final PHPMA polymer was determined by UV-Vis spectroscopy using the calibration curve reported in Figure S3. Taking into account the 19F-PHPMA polymer molecular weight reported in 19 ACS Paragon Plus Environment


Table 1, the degree of chain end functionalization was found to be 80 %. For the studies reported in this paper, a ~ 20 kDa PHPMA polymer was used, as it was previously shown that cellular uptake of these low molecular weight PHPMA polymers is more pronounced as compared to higher molecular weight analogues.16

INSERT SCHEME 1 INSERT TABLE 1

Cell viability and PHPMA internalization Cellular uptake of PHMPA by HeLa cells was assessed using flow cytometry. These experiments, which take advantage of the Rhodamine Red label at the -terminus of the polymer, were carried out at a polymer concentration of 0.6 mg/mL. To assure that at this concentration the viability of the cells is not compromised, the cells were assessed by the Annexin V-Alexa Fluor 647/DAPI assay. For this assay, cells were exposed to polymer for 10 min, 40 min, 1 h, 2 h, 4 h, 15 h or 24 h and subsequently


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incubated with Annexin V-Alexa Fluor 647 and DAPI, which allowed to discriminate between viable, apoptotic and dead cells. While viable cells are not stained by these two dyes, Annexin V-Alexa Fluor 647 binds to the exposed phospholipids of cells undergoing apoptosis. DAPI, which is a membrane impermeable DNA binding dye, stains dead cells that have completely lost membrane integrity. The different regions in the flow cytometry scatter plots that were assigned to viable, apoptotic and dead cells were identified in a separate experiment in which cells were stained with either Annexin V-Alexa Fluor 647 or DAPI (Figure S5). Figure S6 presents scatter plots of 2 independent experiments that were performed to assess the viability of the Hela cells for a period of up to 24 h. The corresponding viabilities, which were determined from these experiments, are presented in Figure 1 and are around 79 % independent of the incubation time.

INSERT FIGURE 1

Uptake of the -19F, -Rhodamine Red labeled PHPMA polymer by Hela cells over a period of 24 hours was followed by flow cytometry. As an example, Figure 2A shows the observed shift in the Rhodamine Red associated fluorescence with increasing incubation time. The result from a duplicate experiment is reported in Figure S7. Figure 2B summarizes the results from these two experiments and represents the



Rhodamine-geometric mean fluorescence intensity (normalized to the geometric mean fluorescence of untreated cells) as a function of time. Figure 2B shows a continuous, steady increase in fluorescence intensity in the first stage of the experiment. After longer incubation times (between 15 h and 24 h), however, the rate of internalization, which can be estimated from the slope of the line that connects the data points in Figure 2B, decreases. This behavior, previously observed for nanoparticles, has been attributed to cell division rather than saturation of cellular uptake. This effect, however, has not been investigated for linear polymers such as PHPMA.25,42 In order to investigate whether the apparent reduction in the rate of polymer uptake at long incubation times can also be (partly) attributed to cell division, a CellTrace violet proliferation assay was performed. CellTrace Violet diffuses into the cell and binds covalently to intracellular amines. Hence, a timedependent decrease in CellTrace Violet associated fluorescence in, for example, a flow cytometry experiment can be used as an indication of cell division. For this experiment, cells were treated using the same conditions as for the polymer uptake study and flow cytometry was used to detect CellTrace Violet associated fluorescence. Figure 3A illustrates the design of this experiment and Figure 3B 22 ACS Paragon Plus Environment

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summarizes the CellTrace Violet assay results. The corresponding flow cytometry plots and geometric mean fluorescence values are reported in Figure S8. For all experiments, cells were seeded 18 hours before starting the time course polymer incubation (“seeding time”). For each incubation time, one sample of cells was analyzed by flow cytometry in order to determine the CellTrace violet associated fluorescence at the beginning of the time-course experiment. The fluorescence of cells analyzed at the end of the experiment is indicated in Figure 3 as “End time”. The shift between “Seeding time” and “End Time” fluorescence suggests that during this time frame, cells underwent at least two division cycles. The geometric mean fluorescence of cells analyzed for the time points “10 min”, “40 min” “2 h” and “4 h” overlaps with that of the cells at the end of the time-course experiment “End time”. This result indicates that both cell division cycles occurred prior to polymer addition. On the contrary, for the time points “15 h” and “24 h”, cell fluorescence values prior to polymer addition were found in between the “Seeding time” and “End time” values. This suggests that, in this case, cells underwent at least one division cycle during the polymer incubation period. As a consequence, the apparent decrease in the rate of cellular uptake of the PHPMA polymer that is indicated in Figure 2B may be 23 ACS Paragon Plus Environment


attributed both to saturation effects as well as cell division during the time course of the experiment.

INSERT FIGURE 2 INSERT FIGURE 3

Intracellular trafficking of PHPMA via confocal microscopy

To investigate the intracellular trafficking of the PHPMA polymer, co-localization studies were performed with Hela cells that express GFP either in the early endosomes (EE) or lysosomes (L). For these experiments, Hela cells expressing GFP either in the early endosomes or in the lysosomes were incubated with the polymer for 24 h, 15 h, 4 h, 2 h, 40 min and 10 min and subsequently fixed and imaged by confocal microscopy. Figure 4 shows representative images of EE or L GFP expressing HeLa cells taken 10 min or 4 h after incubation with the polymer. In each image, panels a and d represent the threshold images corresponding to endosomal or lysosomal vesicles (panel b, in green) and polymer (e, in red), which were determined imaging control samples stained only with GFP or Rhodamine. The fluorogram as well as the “co-localized” pixels are also reported in panels c and f, respectively. While 10 minutes incubation gave rise 24 ACS Paragon Plus Environment

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to very little polymer accumulation in cells (low Rhodamine intensity in Figure 4A and 4C), 4 h incubation already resulted in an increased signal intensity (Figure 4B and 4D). Endosomal co-localization was found to be higher in the 10 minutes incubation (Figure 4A, panel f) rather than longer incubation time such as 4 h (Figure 4B, panel f), on the contrary, predominant lysosomal colocalization can be observed at 4 h incubation as illustrated in Figure 4D panel f. This indicates that with increasing incubation times, polymer continues to traffic down the endocytic pathway from the early endosomes to the lysosomes. Longer incubation times resulted in even higher intensities of the polymer-associated signal as shown in Supporting Information Figure S9 and Figure S10.

INSERT FIGURE 4

Two different image analysis methods were employed to process the confocal microscopy images and investigate the kinetics of the endocytosis and to determine the time dependent polymer co-localization with both endosomal and lysosomal vesicles. The first analysis investigates the change in area ratio between endolysosomal vesicles (GFP, Channel 1) or Rhodamine (19F-PHPMA-Rhodamine, Channel 2), with respect to the area of the Regions of Interest (ROIs) as a function of incubation time. Figure 5 presents the area ratio plots. Figure 5A and Figure B plot the



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“Area Threshold in Channel 1 (Ch1 GFP) normalized by “the area of the ROI” (ROIs Area), respectively, “Area Threshold in Channel 2 (Ch2 Rhodamine) normalized by “the area of the ROI” (ROIs Area) for Hela cells with GFP expressing early endosomes. Figure 5C (Ch1, GFP) and Figure 5D (Ch2 Rhodamine) present the same plots for HeLa cells with GFP expressing lysosomes. For each time point, Figure 5A – 5D show datapoints that were obtained from the analysis of images from three independent experiments.

INSERT FIGURE 5

Image analysis revealed that for both early endosomes (Figure 5A) and lysosomes (Figure 5C), the area of Ch1 (GFP, EE or L) was found to slightly increase over time, especially at the latest time points, i.e., 15 h and 24 h. This effect was more pronounced in the case of lysosomal staining (Figure 5C).43 The continuous increase in Ch1/ROI in Figure 5A and Figure 5C indicates an enhanced formation of EE and L upon increasing the exposure time of the cells to the polymer.43 This observation was confirmed by transmission electron microscopy (TEM) analyses of the cells. Figure S11 shows representative images of control cells as well as cells treated with the polymer for 4 and 15 h. In these images, endolysosomal vesicles are visible as dark stained organelles. The images illustrate that cells exposed to the polymer solution contain a higher number of endolysosomal vesicles as compared to control cells. Analysis of the evolution of the Ch2/ROI ratio with time (Figure 5B and Figure 5D) indicates that for both the EE and L the enhanced formation of these intracellular compartments correlates with enhanced uptake of the 26 ACS Paragon Plus Environment

19F-labelled

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PHPMA polymer. This increased cellular internalization is in agreement with the flow cytometry results (Figure 2).

While the first analysis method illustrates the continuous polymer uptake during the investigated time, a second image analysis technique was used to study the colocalization of the

19F-PHPMA-Rhodamine

polymer and endolysosomal vesicles (EE

and L) and to determine the intracellular fate of the internalized polymer at different incubation times. To this purpose, Mander’s coefficients were determined.44 These coefficients are proportional to the amount of fluorescence of the co-localizing pixels in each channel. Values ranging from 0 to 1 express the fraction of intensity in a channel that is located in pixels where there is above zero (or threshold) intensity in the other channel.35 Hence, in this case, for Ch1 (GFP, EE or L) and for Ch2 (Rhodamine,

19F-PHPMA-Rhodamine),

M1 and M2, respectively, were determined.

The equations for these coefficients are expressed in Equations (1) and (2) in the Supporting Information.35

Figure 6 presents Mander’s Coefficient plots for cells with GFP expressing early endosomes or lysosomes and provides information about the colocalization between the polymer and 27 ACS Paragon Plus Environment


endolysosomal vesicles for different incubation times. For each time point, Figure 6A – 6D show datapoints that were obtained from the analysis of images from three independent experiments. Figure 6A presents the Mander’s coefficients determined for images of cells expressing the endosomal marker (EE). At the earliest time point “10 min”, the M1 values are very low (all below 0.25, mostly below 0.125), indicating little co-localization of Ch1 intensities (GFP, EE) with the Ch2 (Rhodamine, 19F-PHPMA-Rhodamine). This suggests that only a small fraction of the EE vesicles are loaded with the polymer. On the other hand, M2 values were observed, which range from 0 to 1 (Figure 6B), indicating that some cells showed little, while other a high degree of co-localization of Ch2 (Rhodamine, 19F-PHPMARhodamine) intensities with Ch1 (GFP, EE). These results suggest that at the earliest time point, the polymer already entered the early endosomes, but only a very small fraction of these vesicles are occupied (M1 values are low). At “40 min” no change for the M1 values is found while the M2 values decrease remarkably. At the 2 h time point, the M2 values for some of the examined images further decreases. Time points 4 h and 15 h shows a more variable cell population but always characterized by increasing M1 values. Hence, after the first time point, while the M1 values slightly increase, suggesting that more EE are involved in the endocytosis process, the M2 values decreases, indicating that more and more of the internalized polymer is not localized inside the endosomal vesicles. This result, together with the increase of the Ch2 area ratio over time (Figure 5B) suggests that the internalized polymer chains, first detected in the endosomal vesicles, continue their route to other compartments, most probably the lysosomes. In order to prove this hypothesis, the same analysis was performed for lysosomal co-localization. Mander’s coefficients analysis (Figure 6C) revealed that at the earliest time point 10 min, the M1 values are very low (mostly below 0.25), suggesting little co-localization of Ch1 intensities with the Ch2. On the other hand, the M2 values are mainly above 0.5 (Figure 6D). These results suggest that even at the earliest 28 ACS Paragon Plus Environment

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time point, some polymer has already reached some lysosomes (M2 values), but just a few of the lysosomes are involved in the process (M1 values are low). At “40 min”, both M1 and M2 increase and values are more homogenous. This suggests that more polymer has trafficked to the lysosomes, and more lysosomes have been occupied. At the subsequent time points, i.e. between “2 h” and “15 h”, M1 values increase gradually, suggesting that more and more lysosomes are taking part in the process.

INSERT FIGURE 6

The confocal microscopy experiments, and in particular the image analysis discussed above, confirmed the rapid and continuous polymer uptake previously observed via flow cytometry. Moreover, it was also observed that during the first 10 min of incubation, polymer was already detected in the lysosomes. After 40 min incubation, the polymer detected in the early endosomes drastically decreased, while co-localization with lysosomal vesicles increased and prolonged incubation resulted in continued polymer internalization via the early endocytic vesicles. However, the internalized material was found to only transit through these vesicles to finally accumulate in the lysosomes. Co-localization studies revealed that 2 – 4 h cell exposure to the polymer allows dominant localization in lysosomes and longer incubation times resulted in higher amount of lysosomal vesicles involved in the process. Understanding the time-dependent accumulation of the polymer in the lysosomes is particularly relevant since the environmental parameters that characterize these vesicles can be exploited to trigger intracellular drug release from PHPMA carriers.45 As such the insights obtained from the experiments discussed above may be valuable in the design of polymer 29 ACS Paragon Plus Environment


conjugates or polymer particles that contain linkers that are cleavable e.g. in the lysosomal or endosomal compartments.

Subcellular polymer localization via NanoSIMS and TEM After having characterized the kinetics of the internalization and intracellular trafficking of the PHPMA polymer and knowing the time needed for the polymer to accumulate in the lysosomal compartments, the final aim of this study was to explore the feasibility of NanoSIMS in combination with TEM to localize the polymer in these compartments. As mentioned earlier, NanoSIMS has been previously successfully used to probe the cellular internalization of polymeric nanoparticles.33 In this previous work, however, cells were dehydrated and dry cells were investigated with NanoSIMS. While this allowed to track the internalized carrier and drug, the use of this sample preparation protocol has a number of limitations. First of all, dehydration of the cells may compromise the integrity of the organelles and also does not allow sample sectioning, therefore making it challenging to image and analyze deeper cytoplasmic regions. Another consequence of this sample preparation protocol is that organelles need to be fluorescence stained and fluorescence microscopy is


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needed to map the organelles. For the present study, an alternative sample preparation protocol was explored, which involved embedding of the cells in resin. In this way, a drying out of the cells could be avoided. This is advantageous as the organelles can be visualized with TEM and NanoSIMS be used to directly monitor the intracellular localization and distribution of the polymer. For this proof-of-concept study, cells were investigated that had been exposed to the PHPMA polymer for 4 or 15 h. These time points were selected based on the results of the confocal microscopy image analyses, which revealed a preponderant polymer co-localization with the lysosomes after exposure of cells for 4 hours or more to the polymer, as discussed above. Untreated cells were used as a control. After confocal microscopy, the samples were heavy metal stained, embedded in resin, sectioned at 70 nm thickness and imaged by TEM. Representative images are shown in Figure S11. As mentioned earlier, exposure of cells to the polymer resulted in an increase in the number of lysosomal vesicles as compared to control cells that were not exposed to polymer.



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Next, resin embedded samples were analyzed by NanoSIMS. Examples of secondary ion maps of control cells, as well as of cells incubated with the polymer for 4 h and 15 h are shown in Figure 7. Figure S12 presents secondary ion maps of other selected cells from the same experiment. These results show that fluorine-containing polymers can be detected intracellularly as punctate signals and that longer incubation times result in an increase in the number and intensity of these intracellular fluorine signals. Interestingly, Figure 7 indicates that the intracellular fluorine signal strongly correlates with sulphur. Localized sulphur in the cell cytoplasm has been previously observed by NanoSIMS and has been attributed to sulphur-rich lysosomal vesicles.32 As the polymer used in this study also contains sulphur, the signals in the

32S-

ion maps of cells exposed to the polymer are most likely due to both the

sulphur rich lysosomal vesicles as well as the polymer. The punctuated 19F- signals in Figure 7B and Figure 7C are due to the 19F label that is present in the polymer only. These results indicate that NanoSIMS 19F- secondary ion imaging can be used for co-localization studies of polymers with lysosomal vesicles, in principle without the need for other imaging techniques.

INSERT FIGURE 7

To unambiguously confirm the co-localization between the PHPMA polymer and the endolysosomal vesicles, selected cells were imaged first by TEM and subsequently analyzed by NanoSIMS. Figure 8 shows a TEM image and

19F

and

32S

secondary

ion maps from the same cell, which illustrate the correlation between these two techniques. Figure S13 presents a TEM image and secondary ion maps of another 32 ACS Paragon Plus Environment

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selected cell from the same experiment. As expected, fluorine signals correlating with sulphur were co-localized with electron dense endolysosomal vesicles. This result confirms the potential of NanoSIMS to monitor polymer accumulation in cell lysosomes and suggests that correlation with TEM images is a powerful approach to visualize intracellular cargo distribution.

INSERT FIGURE 8

CONCLUSIONS The aim of this study was to explore the feasibility of NanoSIMS as a fluorescentlabel free technique to study the intracellular distribution of polymer – drug conjugates. For the proof-of-concept study reported here, as a prototypical polymer – drug conjugate, a PHPMA polymer was used, which contained a

19F-label

for

NanoSIMS analysis at the -terminus as well as a Rhodamine-dye label at the other chain end. The fluorescent label was introduced to study the internalization of the polymer by confocal microscopy and flow cytometry. These studies provided detailed 33 ACS Paragon Plus Environment


insight into the internalization kinetics of the polymer and were used to select the incubation times for the NanoSIMS experiments. For the experiments reported here, cells were investigated that had been exposed to polymer for incubation times that lead to predominant localization in the lysosomal compartments. Cells for NanoSIMS analysis were resin-embedded and sectioned into 70 nm thick samples, which also facilitated simultaneous TEM analysis. For HeLa cells exposed to the PHPMA polymer for 4 or 15 hr, the NanoSIMS secondary ion maps revealed a punctuate intracellular fluorine signal that was found to strongly correlate with a sulphur signal that was attributed to sulphur-rich lysosomal vesicles. TEM analysis of exactly the same cell sample allowed to correlate the location of the NanoSIMS fluorine and sulphur signals with the position of the endolysosomal compartments. The results of this study provide a first proof-of-concept that demonstrates the feasibility of NanoSIMS to monitor the intracellular distribution of polymer conjugates without the need for a fluorescent label. As NanoSIMS allows the parallel acquisition of multiple masses, this technique may also be of use to track intracellular drug release and monitor independently the trafficking and fate of polymer carriers and release drugs.


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SUPPORTING INFORMATION Supporting Information is available free of charge on the ACS Publications website at DOI: XXXXXXX. Equations for the calculation of the Mander’s coefficients, NMR spectra of the polymer, calibration curve for the determination of the degree of Rhodamine end functionalization, additional flow cytometry, confocal microscopy as well as TEM images and NanoSIMS secondary ion maps.

ACKNOWLEDGMENTS This research has been financially supported by the Swiss National Science Foundation (SNSF).

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Table 1. Molecular weights and dispersities (Mw/Mn) of the 19F-PPFMA and 19FPHPMA polymers investigated in this study.

MnTH,a

MnNMR,b

MnSEC

Mnc

Mw/Mn

(g/mol)

(g/mol)

(g/mol)

(g/mol)

(-)

19F-PPFMA

21180

33000

30200

-

1.4

19F-PHPMA

-

-

17100

-

Polymer

a

Theoretical number-average molecular weight (MnTH) calculated from the

conversion determined by 19F-NMR spectroscopy of the crude product. b

Number-average molecular weight (Mn) calculated from the

19F-NMR

spectrum of

the purified product. c

Number-average molecular weight (Mn) calculated from the

determined by SEC.


19F-PPFMA

Mn


Scheme 1. Synthesis of the -19F, -Rhodamine-labelled PHPMA polymer used in this study.


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Figure 1. Viability of HeLa cells after incubation with 0.6 mg/mL

19F-PHPMA-

Rhodamine for different times as determined using Annexin V-Alexa Fluor 647/DAPI assay with flow cytometry. Results are reported as percentage of viable cell population determined from two independent experiments, error bars represent the standard deviation.



Figure 2. (A) Flow cytometry histogram of Hela cells upon incubation with

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19F-PHPMA-

Rhodamine (0.6 mg/mL) at different incubation times. (B) Evolution of the Rhodamine Red associated fluorescence as measured by flow cytometry upon incubation of HeLa cells with

19F-PHPMA-Rhodamine

for 10 min, 40 min, 2 h, 4 h, 15 h and 24 h.

Results are reported as average geometric mean fluorescence determined from two independent experiments. Error bars represent the standard deviation and geometric mean fluorescence has been normalized to the fluorescence of untreated cells.


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Biomacromolecules

Figure 3. (A) Scheme of the CellTrace Violet proliferation assay. (B) Shift of CellTrace Violet cell fluorescence deriving from cell division during the time frame of the experiment as determined by flow cytometry. The grey curve represents the initial cell population (Seeding time), while the final population is indicated in violet (End time). The curves labeled with the different time points represent cell fluorescence prior to polymer addition for each incubation time.



Figure 4. Co-localization studies of

19F-PHPMA-Rhodamine,

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in red, with (A and B) early

endosomes expressing GFP and (C and D) lysosomes expressing GFP, both indicated in green. Representative panels of observations at (A and C) 10 min and (B and D) 4 h

19F-

PHPMA-Rhodamine incubation (0.6 mg/mL) show the threshold images (a and d), corresponding to endosomal or lysosomal vesicles (b, in green) and polymer (e, in red), represented together with the DAPI channel (b, e and f, in cyan). The merged channels with the “co-localized” pixel (f) as well as the fluorogram (c) are also reported. The contours of the manually drawn ROIs are shown as a white line.


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Biomacromolecules

Figure 5. Area Ratio plots for cells with Early Endosomes (EE) and Lysosomes (L) expressing GFP as a function of incubation time. (A) “Area Threshold in Channel 1” (Ch1, GFP) normalized by the “Area of the ROI” and (B) “Area Threshold in Channel 2 (Ch2, Rhodamine) normalized by the “Area of the ROI” for cells with GPF expressing Early Endosomes; (C) “Area Threshold in Channel 1” (Ch1, GFP) normalized by the “Area of the ROI” and (D) “Area Threshold in Channel 2” (Ch2, Rhodamine) normalized by the “Area of the ROI” for cells with GFP expressing lysosomes. For each time point, Figure 5A – 5D show datapoints that were obtained from the analysis of images from three independent experiments.



Figure 6. Mander’s coefficient plots for cells with Early Endosomes (EE) and Lysosomes (L) expressing GFP as a function of polymer incubation time. (A) Mander’s Coefficients of Channel 1 (M1, GFP) and (B) Mander’s coefficients of Channel 2 (M2, Rhodamine) for cells with GPF expressing Early Endosomes. (C) Mander’s Coefficients of Channel 1 (M1, GFP) and (D) Mander’s coefficients of Channel 2 (M2, Rhodamine) for cells with GFP expressing Lysosomes. For each time point, Figure 6A – 6D show datapoints that were obtained from the analysis of images from three independent experiments. Errors bars represent standard deviation.


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Biomacromolecules

Figure 7. Secondary ion maps of

14N12C-, 19F-

and

32S-

in (A) untreated Hela cells (control

cells) and in Hela cells treated with 0.6 mg/mL 19F-PHPMA-Rhodamine for 4 hours (B) and 15 hours (C).



Figure 8. Representative TEM image and the corresponding

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19F, 32S

and

12C14N

ion maps obtained after 15 h incubation of Hela cells with 0.6 mg/mL

secondary

19F-PHPMA-

Rhodamine. White arrows indicate some of the observed correlations between cell vesicles, 19F-

and 32S- signals, as an example.


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Cellular Uptake and Intracellular Trafficking of Poly(N-(2

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