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HEMATOPOIESIS AND STEM CELLS
“Microenvironmental contaminations” induced by fluorescent lipophilic dyes
used for noninvasive in vitro and in vivo cell tracking
Francois Lassailly,1 Emmanuel Griessinger,1 and Dominique Bonnet1
1Hematopoietic
Stem Cell Laboratory, London Research Institute, Cancer Research UK, London, United Kingdom
Determining how normal and leukemic
stem cells behave in vivo, in a dynamic
and noninvasive way, remains a major
challenge. Most optical tracking technologies rely on the use of fluorescent or
bioluminescent reporter genes, which
need to be stably expressed in the cells of
interest. Because gene transfer in primary leukemia samples represents a major risk to impair their capability to engraft in a xenogenic context, we evaluated
the possibility to use gene transfer–free
labeling technologies. The lipophilic dye
3,3,3ⴕ,3ⴕ tetramethylindotricarbocyanine
iodide (DiR) was selected among 4 nearinfrared (NIR) staining technologies. Unfortunately we report here a massive
transfer of the dye occurring toward the
neighbor cells both in vivo and in vitro.
We further demonstrate that all lipophilic
dyes tested in this study (1,1ⴕ-dioctadecyl3,3,3ⴕ,3ⴕ-tetramethylindotricarbocyanine perchlorate [DiI], DiD, DiR, and
PKH26) can give rise to microenvironmental contamination, including when
used in suboptimal concentration, after
extensive washing procedures and in
the absence of phagocytosis or marked
cell death. This was observed from all
cell types tested. Eventually, we show
that this microenvironmental contamination is mediated by both direct cell-cell
contacts and diffusible microparticles.
We conclude that tracking of labeled
cells using non–genetically encoded
markers should always be accompanied
by drastic cross validation using multimodality approaches. (Blood. 2010;115(26):
5347-5354)
Introduction
Despite recent animated discussions,1 xenotransplantation of human cells in immunodeficient animals remains the only available
technique to analyze the “stemness” of human cancer-initiating
cells.2-5 Despite the wide use of this model, homing and dynamic
behavior of injected cells depending on time remain largely
unknown both for normal and malignant cells. Whole-body imaging (WBI) and intravital microscopy (IVM) provide powerful tools
to longitudinally monitor the behavior of cancers in vivo6,7 and to
track and analyze normal and cancer-initiating cells in their
microenvironment or niche.8,9 Near-infrared optical imaging (NIR)
represents a very attractive technology in this context by offering a
relatively easy and cheap access to multimodality and multiscale in
vivo imaging.10,11
In vivo cell tracking using optical imaging technologies
largely relies on the use of ectopic expression of reporter genes
such as Luciferase (for bioluminescence) or on fluorescent
proteins. To obtain such expression, cells have to be engineered
in vitro so that the reporter gene can be integrated in their
genome for stable expression over time. Because in vitro
manipulation of primary leukemia samples represents a real risk
to impede the functionality of the cells, gene transfer–free
approaches would represent a major improvement in this
context.12 Various strategies have been reported to fluorescently
label cells in vitro for subsequent tracking in vivo.13-16 In the
field of immunohematology, the succinimidyl ester of carboxyfluorescein diacetate (5(6)-CFDA SE), or the closely related but
distinct carboxyfluorescein succinimidyl ester (CFSE) as well as
PKH26 are the gold standard techniques for cell tracking. Gene
transfer–free labeling of cells relies mainly on 3 technologies.
Amine-reactive probes (such as CFSE) diffuse passively into
cells where they will react with cytosolic amine-containing
residues to form dye-protein adducts that are retained in cells.
We have tested here a far-red derivative of this technology is the
DDAO-SE. Lipophilic dyes (such as PKH26) are based on
membrane labeling by stable incorporation of long-chain carbocyanine dye containing long aliphatic tails into lipid regions of
the cell membrane.17 A wide variety of these molecules have
been developed in terms of spectral properties, including in the
far-red and NIR windows, which make them attractive candidates for WBI and confocal IVM (DiD or 3,3,3⬘,3⬘ tetramethylindotricarbocyanine iodide [DiR]). Nanoparticles are the third
type of technology usable for cell labeling. Fluorescent quantum
dots are small nanocrystals (1-10 nm) made of cadmium, an
inorganic semiconductor material, possessing several unique
optical properties well suited for in vivo imaging. The dots are
very photo stable, remain resistant for long periods of time in
vivo, and are available in a large variety of colors, including in
the NIR region (quantum dot 705 nm). In addition, their
excitation spectrum makes them ideal tools for 2-photon
excitation.18 Eventually, we also evaluated a fourth family of
strategies consisting of using an antibody labeling kit (IRDye
800CW) to covalently stain the extracellular domain of membrane proteins. As previous studies have nicely shown that
carbocyanine lipophilic dyes are very effective tools for WBI
and/or IVM tracking of normal and malignant bone marrow
(BM)–derived cells without affecting their functionality,19-23 we
aimed at using similar strategies to track primary human acute
myeloid leukemia (AML) and hematopoietic stem cells (HSCs).
Submitted May 26, 2009; accepted February 17, 2010. Prepublished online as
Blood First Edition paper, March 9, 2010; DOI 10.1182/blood-2009-05-224030.
The publication costs of this article were defrayed in part by page charge
payment. Therefore, and solely to indicate this fact, this article is hereby
marked ‘‘advertisement’’ in accordance with 18 USC section 1734.
The online version of this article contains a data supplement.
© 2010 by The American Society of Hematology
BLOOD, 1 JULY 2010 䡠 VOLUME 115, NUMBER 26
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LASSAILLY et al
Methods
Human and mouse cells
Cord blood cells were obtained under research ethics committee approval
and informed consent at the Royal London Hospital in accordance with the
Declaration of Helsinki. Mononuclear cells were obtained by density
centrifugation.
The human HL-60 cell line was provided by the cell culture service at
the Institute and cultivated in RPMI supplemented with 10% fetal calf
serum (FCS), L-glutamine, and penicillin/streptomycin. Transduction of
HL-60 cells was performed with an HIV-1–based self-inactivating (SIN)
lentiviral vector (pHR⬘SINcPPT-SEW-SffV-IRES-eGFP lentivirus), which
carries the enhanced green fluorescent protein (eGFP) reporter gene under
the control of the spleen focus-forming virus LTR (a kind gift from Prof A.
Thrasher, Institute of Child Health, London, United Kingdom) as previously described.24
The human osteosarcoma cell line Cal-72 (purchased from German
Collection of Microorganisms and Cell Cultures [DSMZ], http://
www.dsmz.de) and maintained in Dulbecco modified Eagle medium with
10% FCS ⫹ 2mM L-glutamine ⫹ 1⫻ insulin-transferrin-sodium selenite.
Human umbilical vein endothelial cells (HUVECs; Clonetics) were cultured in endothelial growth medium 2 (Clonetics), and GP293T cells (kind
gift from Prof Eric So, Institute of Cancer Research, London, United
Kingdom) were maintained in Dulbecco modified Eagle medium supplemented with 10% FCS and 1% antibiotics.
Mouse hematopoietic stem and progenitor cell staining and
fluorescence-activated cell sorting
Bone marrow from either ROSA26-eGFP (Biological Resource Unit at Clare
Hall Institute, London, United Kingdom) or ␤-actin–Luciferase (XenogenCaliper) mice was obtained by crushing the bones with a mortar and a pestle in
PBS supplemented with 2% FCS, and cells were harvested after Ficoll gradient
centrifugation and stained using anti–Sca-1–phycoerythrin (PE)–cyanin 7, CD117–
allophycocyanin (APC), and lineage (Lin)–biotinylated (subsequently labeled
with PE-conjugated streptavidin)–specific monoclonal antibodies (see antibody
list in supplemental Methods, available on the Blood Web site; see the Supplemental Materials link at the top of the online article) following the manufacturer’s
instructions. Dead cells (DAPI⫹ [4,6 diamidino-2-phenylindole–positive]) and
doublets (displaying side scatter width) were excluded. EGFP⫹ mature cells
(Lin⫹), early progenitors (Lin⫺ Sca⫺ Kit⫹; LK), and stem-enriched cells (Lin⫺
Sca⫹ Kit⫹) were sorted after exclusion of dead cells (DAPI⫹) and doublets using
a MoFlow cytometer (Beckman Coulter) equipped with a 70-␮m nozzle.
BLOOD, 1 JULY 2010 䡠 VOLUME 115, NUMBER 26
Recovery of the bone marrow of mice that underwent
transplantation for flow cytometric analysis
Animals were killed and bones (femur, tibia, helium) and spleen were
collected. The bone marrow was harvested either by flushing long bones25
or by crushing the bone as described in “Mouse hematopoietic stem and
progenitor cell staining heading.” The cells were filtrated on a 70-␮m cell
strainer, and leukocytes were recovered after red cells underwent ammonium chloride lysis.
In vitro analysis of biocompatibility
Competitive proliferation assay using HL60 cells. Untransduced HL60
cells (HL60-eGFP⫺) were stained with different NIR dyes at different
doses. After repeated washing and incubations, cells were mixed together
with 30% of unstained HL60-eGFP⫹ cells and cultured in standard
conditions. At different times points, cells were harvested and analyzed by
flow cytometry. The principle of this “competitive proliferation assay” is
based on the respective proliferation kinetics of NIR-stained (eGFP⫺) and
NIR-unstained (eGFP⫹) HL60 cells. In the absence of any toxic or
cytostatic effect associated with the dye, the relative proportion of both
subpopulations should remain constant. In the case of toxicity related to
NIR treatment, the amount of cells initially stained with a NIR dye should
decrease.
Optimization of staining procedures on selected murine progenitors.
Lineage (lin)–negative cells from ␤-actin–Luciferase mice bone marrow
cells were obtained using the mouse hematopoietic progenitor cell enrichment kit (StemCell Technologies) following the manufacturer’s instructions. After fluorescent staining of the cells, they were plated in serum-free
medium supplemented with optimized cytokine cocktail24 either in 24-well
plates for flow cytometric (FC) analysis of viability and fluorescence
intensity on day 1 or in triplicates in a black 96-well plate for bioluminescence imaging on day 7.
In vitro bioluminescence imaging
Just before imaging, 100 ␮L of luciferin 2⫻ (300␮g/mL; Xenogen-Caliper)
was added to the 100 ␮L of cell suspension in each well. After 5-minute
incubation, bioluminescence activity was acquired in a prewarmed (37°C)
IVIS Spectrum (Xenogen-Caliper) using a 3-second exposure, binning 16
over a total period of 15 minutes. Data were processed using the Living
Image 3.1 software (Xenogen-Caliper) after drawing regions of interests of
identical size (duplications of the region of interest) around individual
wells. Bioluminescence imaging activity was quantified as a flux (photon
per second). Mean and standard values were calculated for each condition
and expressed as a percentage of the respective control.
Vital fluorescent dyes for cell tracking
CellTrace CFSE Cell Proliferation Kit (CFSE), 1,1⬘-dioctadecyl-3,3,3⬘,3⬘tetramethylindocarbocyanine perchlorate (DiI; DiIC18(3); DiI), CellTrace
Far Red DDAO-SE (DDAO), Q-tracker 705 Cell Labeling Kits (Q-tracker
705), Vybrant DiD cell-labeling solution (DiD), and the long-chain
lipophylic 1,1⬘-dioctadecyl-3,3,3⬘,3⬘ tetramethylindotricarbocyanine iodide
(DiR; DiIC18(7)) were purchased from Molecular Probes; PKH26 Red
Fluorescent Cell Linker Kit (PKH) was purchased from Sigma-Aldrich; and
IRDye 800CW protein labeling kit was purchased from Li-Cor Biosciences;
all products were used following the manufacturer’s instructions.
Adoptive transfer of human HL-60 cells in immunodeficient
mice
All animal experiments were performed in compliance with Home Office
and Cancer Research UK (CRUK) guidelines. Nonobese diabetic/severe
combined immunodeficient (NOD/SCID) mice were originally obtained
from Dr Leonard Schultz (The Jackson Laboratory) and bred at Charles
River Laboratories. They were kept in microisolators and fed with sterile
food and acidified water. Mice aged 8 to 12 weeks were irradiated at
375 cGy (137Cs source) 24 hours before intravenous injection of 5 to
10 ⫻ 106 HL-60 cells.
In vitro near-infrared scanning
Near-infrared scanning was performed using the Odyssey infrared imaging
system (Ly-Cor Biosciences). DDAO-SE and Q-tracker 705 nm were
imaged using the 700-nm channel; DiR and IRDye 800CW fluorescence,
using the 800-nm channel.
In vivo near-infrared scanning
Near-infrared scanning was performed using the Odyssey infrared imaging
system (Ly-Cor Biosciences) as described for in vitro scanning. At 12 hours
to 3 weeks after injection of the leukemic cells, animals were killed by CO2
exposure, and shaved by first using a clipper and then hair remover cream.
Matched controls (age- and sex-matched animals) were imaged on the same
scan to provide reliable and objective representation of the background
and/or autofluorescence signals. Postacquisition image processing was
realized using the Odyssey MousePOD Module (Version 2.1) software. To
determine what a specific signal on a test animal was, the same anatomic
location was analyzed on a proper negative control animal. Image displays
were set up so that fluorescence did not come from the control region of
interest.
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BLOOD, 1 JULY 2010 䡠 VOLUME 115, NUMBER 26
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5349
Figure 1. DiR allows sensitive NIR whole-body imaging tracking of human leukemic cells and analysis of
their homing in immunocompromised recipients.
HL60 cells (107) stained with 20␮M DiR (HL60-DiR) were
injected in 4 animals, whereas 2 controls were injected
with unlabeled cells. At 24 hours after injection, 1 control
and 2 animals injected with HL60-DiR were killed and
prepared for whole-body imaging. Frontal acquisition (top
left panel) and dorsal acquisition (top right panel) are
displayed. The same operations have been realized
13 days after the injection (lower panels). The intensity
settings for the acquisition and the processing of the
images are identical for the 4 images. Fluorescence
intensity scale is displayed on the right side of the
images. J indicates jaws; St, sternum; E, elbow; L, liver;
W, wrist; LN, lymph node; K, knee; F, foot; C, calvaria; S,
scapula; Sc, spinal column; and Sp, Spleen.
Cocultures of hematopoietic and stromal cells
Confocal imaging
After different staining and washing procedures (detailed in each experiment), HL60 or eGFP-HL60 cells were plated on pre-established confluent
layers of primary human umbilical vein endothelial cells (HUVECs) or a
fibroblastic cell line (GP293T). At the end of the culture, nonadherent and
adherent cells were harvested by trypsynization. All the cells from the same
well were pooled together for subsequent staining with anti–human CD45
(hCD45) and anti-hCD33 and flow cytometric analysis.
Similarly mouse mature progenitors (Lin⫺ Sca⫺ Kit⫺; L3⫺), early
progenitors (Lin⫺ Sca⫺ Kit⫹; LK), and a population enriched in functionally defined stem cells (Lin⫺ Sca⫹ Kit⫹; LSK) were sorted from ␤-actin–
Luciferase transgenic animals (no phenotype described to date26). Cells
were stained using published procedures27,28 that were further optimized for
this study: 2␮M DiI for 10 minutes in pure PBS at 37°C, followed by
2 washing steps with PBS, 2% FCS and subsequent overnight incubation in
StemSpan (StemCell Technologies) serum-free medium supplemented with
previously optimized cytokine cocktail.24 Cells were then washed and
plated on pre-established confluent monolayers of HUVECs or GP293T
cells in collagen-coated CellBIND (Corning) 96-well plates optimized for
high-quality imaging. After overnight incubation, 1.5 ␮L of anti–murine
CD45 (mCD45)–APC was added in each well, and the plate was incubated
for 30 minutes at 37°C before confocal imaging and subsequent harvesting
of the cells for flow cytometry (similar procedures as for HL60 cocultures).
A second plate was analyzed 48 hours after initiation of the coculture.
Cells in suspension. Cells in suspension were imaged using an upright
Zeiss LSM 510 equipped with a pulsed Ti-Sapphire laser tuned at 870 nm
and a 633-nm laser to excite the quantum dots and the eGFP and the NIR
dyes, respectively. Fluorescence was collected using band-pass filters at
500 to 550 nm and 650 to 710 nm.
HSPCs/stroma cocultures. At the end of the coculture, anti–mCD45APC antibodies were added to each well and placed in the 37°C
environmental chamber of an inverted LSM510 Zeiss confocal equipped
with a Plan-Apochromat 20⫻/0.75 numeric aperture Zeiss lens. DiI and
APC were imaged separately (multitrack configuration) by successive
excitations at 543 nm and 633 nm and collection of fluorescence using
560-nm and 650-nm long-pass filters, respectively. A 10-␮m stack was
acquired (2-␮m step) using a zoom 3 magnification. Resulting images were
processed using Imaris 6.3.0 software (Bitplane) for brightness adjustment
and 3-dimensional reconstruction. Individual and merged channels are
displayed for the cocultures of LSK and LK on endothelial cells.
Flow cytometric analysis
Depending on the need, mice and/or human cells were stained using
fluorescently labeled antibodies25(see antibody list in supplemental Methods). Washed cells were suspended in PBS containing DAPI (4,6 diamidino2-phenylindole) and analyzed using a BD Life Science Research (LSRII)
flow cytometer.
Absolute quantification of dye transfer. After in vivo or in vitro
incubation of stained cells, flow cytometry was used to determine the
absolute amount of fluorescence associated with stained versus neighbor
cells. The total amount of fluorescence was calculated by multiplying the
number of positive events by the mean fluorescence intensity of the whole
population. The same operation was applied for stained cells (HL60 or
purified hematopoietic stem and/or progenitor cells [HSPCs], depending on
the experiment) and for neighbor cells (recipient or stromal cells, depending
on the experiment). The respective contribution of each population was then
expressed as a percentage of total fluorescence.
Statistical analysis
Averages, standard deviation, and Student t tests were calculated using
Microsoft Excel 2004 for Mac (Version 11.5.5). P values less than .05 were
considered as significant.
Results
We first characterized 4 different dyes based on 4 distinct families
of vital staining procedures (DDAO-SE, DiR, IRDye 800CW, and
Q-tracker705) to obtain near-infrared (NIR) labeling of human
acute myeloid leukemia (AML) cell line HL60. The DIR dye was
our best candidate in terms of brightness, stability, lack of toxicity,
and compatibility with multimodality imaging (supplemental
Figure 1).
In vivo tracking of DiR-stained human leukemia cells by
NIR-WBI
DiR-stained HL-60 cells injected intravenously in irradiated NOD/
SCID mice gave rise to a strong signal nicely associated with bones
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LASSAILLY et al
BLOOD, 1 JULY 2010 䡠 VOLUME 115, NUMBER 26
Figure 2. In vivo transfer of DiR dye from human
leukemic cells toward mouse cells. (A) Sublethally
irradiated mice were intravenously injected with
20 ⫻ 106 HL60-eGFP–bright cells stained with 5␮M DIR
and preincubated overnight at 37°C before the infusion.
BM cells were analyzed by FC 4 days afterward. (B left
panels) Frequency and analysis of DiR⫹ cells within
human (eGFP⫹) and mouse (eGFP⫺) cells. (Right panels) Detailed phenotype of live cells positive for DiR.
(C) FC quantification of human cell frequencies within the
BM of recipients based on the detection of DiR⫹ or
eGFP⫹ events (n ⫽ 4). (D) Determination of the absolute
contribution of mouse and human cells within the total
DiR signal: for each animal (n ⫽ 4), the absolute contribution of human (eGFP⫹) DiR⫹ cells was calculated by
multiplying the mean fluorescence intensity by the number of events. The same operation was done for mouse
cells (eGFP⫺) and for the total population of DiR⫹ cells.
Results are expressed as a percentage of total DiR
absolute signals.
24 hours after transplantation (Figure 1 and supplemental Figure 2)
with no apparent decay over a 2-week period of time (Figure 1
bottom panels). This was quite surprising, knowing that HL-60
cells proliferate rapidly in vivo.
DiR⫹ cells was 8 times higher than GFP⫹ cells (1.04% ⫾ 0.39%
versus 0.12% ⫾ 0.09%, respectively; P ⫽ .010; Figure 2C). Quantification of absolute DiR signal in mouse and human cells further
revealed that only 15.4% (⫾ 7.2%) of the DiR signal was specific,
that is, coming from human cells (Figure 2D).
Assessing the specificity of the signal
To address the specificity of the signal, transduced HL60-eGFP–
bright sorted cells were stained using 5␮M DiR for 30 minutes in
complete medium following published procedures,19 incubated
overnight in culture and intravenously injected in sublethally
irradiated NOD/SCID mice (Figure 2A). Fluorescence intensity
and cell viability (⬎ 90%) were controlled before the adoptive
transfer (Figure 2A). As expected, 4 days after injection, virtually
all eGFP⫹ human cells displayed a bright NIR fluorescence and the
presence of DiR⫹ cells within mouse cells (eGFP⫺) was quite low
(0.86% ⫾ 0.29% of total BM; Figure 2B left panel). However,
analyzing the same datasets by first gating on the DiR⫹ population
provides a very different view. Surprisingly, eGFP⫹ cells represented only 11.5% (⫾ 2.5%) of DiR⫹ cells (Figure 2B right panel).
The DiR⫹ eGFP⫺ cells were of murine origin (mCD45⫹) and
coexpressed the dendritic marker CD11c, suggesting a phagocytic
process for the dye transfer. Within the total BM, the frequency of
Optimization of the staining procedure to reduce/abrogate the
microenvironmental contamination
Although the viability of injected cells was good, the phenotype of
DiR⫹ murine cells suggested the dye transfer to be mediated by a
phagocytosis process, corroborating and extending recent reports.29,30 We thus tested, using the HL60 cell line, 3 different
lipophilic dyes (DiI, DiR, and PKH) and an amine-reactive dye
(CFSE) at very low doses (2 and 0.5␮M31) to determine whether
we can decrease this transfer. The stained cells were coincubated on
a fibroblast layer unable to support hematopoiesis (GP293T cells)
either after 2 washings or after overnight incubation in liquid
culture. Despite suboptimal staining efficiency with the low dye
concentration used (supplemental Figure 3), a substantial amount
of fluorescence in stromal cells was still detected with the
3 lipophilic dyes (but not with the amine-reactive dye; Figure 3 and
supplemental Figure 3). This transfer was significantly reduced but
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BLOOD, 1 JULY 2010 䡠 VOLUME 115, NUMBER 26
Figure 3. Microenvironment contamination occurs from all lipophylic dyes
tested. HL60 cells were stained in PBS without serum with CFSE (0.5␮M),31 DiI, DiR,
or PKH26 (2␮M) for 10 minutes and washed twice in PBS, 2% FCS. Stained cells
were either directly plated on a confluent layer of fibroblasts (GP293T; condition
0 hours) or incubated for 24 hours in standard culture conditions before being plated
on fibroblasts (condition 24 hours). After 24 hours of coculture at 37°C, cells were
recovered by trypsinization, stained for hCD45 ⫹ hCD33 (both PE in the case of DiR,
APC for all other dyes), and analyzed by flow cytometry. The absolute amount of
fluorescence associated with the fibroblasts is displayed as a percentage of the total
fluorescence for each dye (average and standard deviation obtained from 4 wells;
data are representative of 2 independent experiments). Comparing 0 hours versus
24 hours for each dye, the differences are statistically significant (*P ⬍ .004 for all
dyes except CFSE where P ⫽ .7). After 24-hour coculture, all dye transfers using
lipophilic dyes are higher than with CFSE (**P ⬍ .001).
not abrogated by culturing stained HL60 cells overnight before
starting the incubation with fibroblastic cells (Figure 3).
As our ultimate goal was to work with primary AML cells and
knowing, from our experiments and others,12 that these cells are
very fragile and that leukemia-initiating cells (LICs) are difficult to
maintain in vitro, we thus wondered whether alternative procedures
could be used to lower the dye transfer. First, we tested the
possibility of performing the labeling incubation in serum-free
medium to keep the cells in a suitable environment, but this
abrogated the initial staining as one might have predicted (supplemental Figure 5A). After labeling the cells with DiI, DiD, or DiR,
we assessed the beneficial impact of 2, 3, or 4 washing steps, or
even fluorescence-activated cell sorting (FACS) of the stained cells
before starting the coculture (supplemental Figure 5B-C). None of
these procedures was effective. In our hands, reduction in dye
concentration plus overnight incubation was the best strategy to
reduce the transfer and was used in subsequent experiments.
Can we reduce the microenvironmental contamination using
primary stem and progenitor cells?
At that stage, we had only evidence of the dye transfer using a cell
line. During the course of this work, very impressive studies have
demonstrated that lipophilic dyes are very useful tools for tracking
of different primary cells in the bone marrow of live animals by
intravital microscopy.19,23,27,32,33 We thus wondered whether using
primary cells the transfer could be decreased. In preliminary
experiments, we observed that human cord blood cells stained with
DiR could induce transfer in coculture with osteoblastic cells. The
transfer induced by stained osteoblasts was also demonstrated.
The biocompatibility of the staining procedure was further
optimized on selected murine progenitor cells (supplemental
Figure 6). During this step, we noticed that cell viability was
slightly affected by the 10-minute incubation at 37°C in PBS alone
(used for DiI and DiR staining) and even more using buffer C (used
for PKH staining; supplemental Figure 6A representative of
2 independent experiments) even in the absence of any dye
addition. Based on quantification of viable cell numbers by in vitro
bioluminescence, we found that DiI at low concentration gave the
PITFALLS ASSOCIATED WITH LIPOPHILIC DYES TRANSFER
5351
best results in terms of biocompatibility (supplemental Figure 6C).
The relatively low staining efficiency further encouraged us to use
this condition (supplemental Figure 6B).
To determine whether primary hematopoietic stem and/or
progenitor cells (HSPCs) could be affected by this dye transfer in a
short-period of time, stem cell–enriched early progenitors (Lin⫺
Sca-1⫹ Kit⫹ [LSK]) were purified, stained with 2␮M DiI for
10 minutes, washed, incubated overnight in serum-free medium
supplemented with cytokines,24 and washed again before being
cultivated on a confluent layer of unstained endothelial cells for
18 hours. Live confocal imaging of the cells demonstrated clear
transfer in this very stringent condition (Figure 4B), and absolute
quantification revealed that more than 30% of DiI fluorescence was
associated with endothelial cells (Figure 4C). This result demonstrates that, even in the absence of marked cell death (Figure 4A),
transfer occurs not only with primary cells but also with HSPCs. In
an independent experiment, similar results were obtained when
LSK cells stained with DiR were cultivated on a complex stroma
composed of human osteoblasts, endothelial cells, and murine
stromal cells for 12 hours (data not shown). Although the intensity
of the transfer is affected by the duration of the incubation, overall
substantial amounts of dye transfer occurred during this period of
time (supplemental Figure 4). Furthermore, we wondered whether
we could used this strategy for “tattooing” the stromal cells directly
interacting with HSCs in vivo (stem cell niche), we performed the
same experiment in parallel with more engaged progenitors devoid
of stem cell activity (Lin⫺ Sca-1⫺ Kit⫹ [LK] and Lin⫺ Sca-1⫺ Kit⫺
[L3⫺[). Because endothelial cells might display the capability of
supporting both stem cells and more committed progenitors, the
experiment was conducted in parallel on hematopoietic nonsupportive fibroblastic cells. As expected, hematopoietic cell viability was
slightly reduced after coincubation with fibroblasts compared with
endothelial cells (Figure 4A), illustrating different functionalities
of the 2 stromal cell types used. A transfer could be detected and
quantified in all conditions tested (Figure 4B-C), demonstrating
that this phenomenon is not a specific feature of stem cells, and that
a stem cell niche–type interaction is not necessary for this to occur.
Mechanisms of transfer of the lipophilic dyes
In preliminary experiments, where DiR-stained cord blood mononuclear cells (MNCs) were incubated with osteoblasts, the 2 cell
populations being separated by a microporous membrane, we
observed that the transfer on osteoblasts was only partly reduced,
suggesting that diffusible elements could be involved in the process
and that a direct contact between donor and acceptor populations
was not absolutely required (data not shown).
Because the transfer occurred similarly with different subpopulations of murine HSPCs, human cord blood MNCs, and HL60 cell
line, we chose to further dissect the mechanism of transfer with
HL60 cells considered as a homogeneous population (Figure 5).
HL60 cells stained with DiI were cultivated overnight. Cells were
recovered after centrifugation, and the upper part of the supernatant
was further fractionated by ultracentrifugation to produce the
ultrapellet and the ultrasupernatant. Similar quantities of cells, and
equivalents of ultrapellet and ultrasupernatant, were dispatched on
confluent stromal monolayers and incubated at 37°C. We observed
that both direct cell-cell contact and the ultrapellet gave rise to
transfer. Of interest, no transfer was obtained from the ultrasupernatant fraction. Similar data were obtained using DiR in 2 independent experiments, whereas no transfer could be detected when
eGFP-bright HL60 cells were used. This confirmed our preliminary
observations on cord blood and osteoblastic cells and further
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LASSAILLY et al
Figure 4. Microenvironment contamination occurs
from sorted HSPCs but does not reflect a stem cell
niche–type interaction with stromal cells. Different
populations of HSPCs (Lin⫺ Sca-1⫹ Kit⫹ [LSK], Lin⫺
Sca-1⫺ Kit⫹ [LK], Lin⫺ Sca-1⫺ Kit⫺ [L3⫺]) that were FACS
purified (purity ⬎ 90% for all fractions) were stained with
2␮M DiI, washed, and incubated overnight in serum-free
medium complemented with cytokines. After washing,
cells were plated on confluent layers of endothelial cells
(HUVECs) or fibroblastic cells (GP293). After the coculture, dye transfer was analyzed by confocal imaging (B)
and FC (A,C). (A) Cells were recovered by trypsinization
and stained with anti–mCD45-APC. After washing, cells
were suspended in DAPI containing PBS, 2% FCS for FC
analysis. After elimination of debris and doublets, viability
(DAPI⫺) was determined on HSPCs (CD45⫹). (B) After
overnight coculture (18 hours), hematopoietic cells were
stained in situ by an anti–mCD45-APC antibody. A 10-␮m
stack was acquired for each region of interest (2-␮m step)
using a zoom 3 magnification. Displayed are 3-dimensional reconstructions for individual and merged channels for the cocultures of LSK and LK on endothelial cells.
Scale bar represents 20 ␮m. (C) Absolute quantification
of DiI distribution after 22 hours of coculture: using the
same strategy as in Figure 2D, absolute quantification of
DiI fluorescence was measured for HSPCs (CD45⫹) and
stromal cells (CD45⫺) and expressed as percentage of
total DiI fluorescence in the different conditions.
demonstrated that transfer can occur from diffusible elements that
can be recovered by ultracentrifugation, which implies that it is not
mediated by the release of free dye molecules in the environment.
These results show that lipophilic dye transfer to neighbor cells
most certainly occurs via the exchange of membrane microdomains, a mechanism known as trogocytosis,34 but also probably via
microvesicle and/or exosome35,36 transfer.
Discussion
Figure 5. Microenvironment contamination is mediated by direct cell-cell
interactions and diffusible particles but not free dye molecules. HL60 cells were
stained using 4␮M DiI for 10 minutes in PBS without FCS at 37°C, washed, and
plated in standard liquid culture conditions. As a control, eGFP-bright cells were
treated in identical conditions without staining. Before and after culture, cell viability
was higher than 90% (not shown). After 24-hour culture, cells were centrifuged (300g
for 7 minutes at 4°C). The upper third part of the supernatant was harvested for
ultracentrifugation (compare below); the remaining one was discarded. Cells were
washed and 100 000 cells from each condition were plated on a confluent layer of
HUVECs in a well of a 24-well plate. The supernatant from the first centrifugation step was
further ultracentrifuged (61 600g for 2 hours 30 minutes at 4°C). The resulting supernatant
(ultra-SN) and pellet (ultrapellet) were harvested separately. We determined the amount of
ultra-SN and ultrapellet that would be equivalent to the number of cells plated on HUVECs
and added this amount into separate confluent HUVEC wells. After 24 hours of culture,
hematopoietic and stromal cells were recovered and analyzed by FACS after staining with
anti–human CD45-APC and anti–human CD33-APC monoclonal antibody. DiI contamination of stromal cells (CD45⫺) after direct coculture or after exposure to similar amounts of
ultrapellet or ultra-SN is displayed. No eGFP signal was found associated with stromal
cells, whatever the condition (upper line). These data are representative of 2 experiments,
which also included DiR.
In this study, we have conducted a comprehensive analysis
concerning a variety of vital dyes based on 4 distinct technologies
including amine-reactive dyes (CFSE [green] or DDAO-SE [far
red]), lipophilic dyes (DiI [red], PKH26 [red], DiD [far red] and
DiR[NIR]), nanocrystals (quantum dots 705 [NIR]), and a kit for
labeling antibodies (IRDye 800CW [NIR]). At the end of a
thorough assessment, we were not able to find a single dye that
would fulfill all of our initial criteria of multimodal imaging
combining WBI, IVM, and flow cytometry and also provide good
evidence of reliability. CFSE displayed very good biocompatibility
and staining efficiency, while not displaying microenvironmental
contamination, which make this strategy very suitable to track cells
in different settings. Although the green fluorescence emitted by
CFSE is not suitable for sensitive whole-body imaging,10 our data
confirm it is a very reliable tool for microscopy (including 2-photon
microscopy) and flow cytometric analyses. The infrared version
DDAO-SE is compatible with all imaging techniques tested but
was toxic for the cells in our study. Although quantum dots did not
display major toxicity on HL60 cells, they provided very heterogeneous staining, which is not suitable for IVM imaging. Although
the main limiting factor observed for IRDye 800CW was linked to
From www.bloodjournal.org by guest on December 22, 2014. For personal use only.
BLOOD, 1 JULY 2010 䡠 VOLUME 115, NUMBER 26
its suboptimal excitation by 633-nm lasers used for IVM and
FACS, we did not try to use an antibody labeling kit more
compatible with our imaging systems because it does stain
membrane proteins. Indeed, the transfer of membrane antigen was
reported and might lead to microenvironmental contamination
(MEC) in a similar manner as what we describe for lipophilic
dyes.34,37,38
To our knowledge, this study is the first to demonstrate that
lipophilic dyes (DiI, PKH-26, DiD, and DiR) used to stain human
leukemic cells, cord blood MNCs, but also different sorted
populations of murine HSPCs, can rapidly induce (within 1524 hours) MEC in the absence of phagocytosis (Figures 3-4 and
supplemental Figures 3-4). Fluorescent MEC could not be detected
using amine-reactive CFSE or eGFP-encoded reporter vector,
excluding a general mechanism involving all types of fluorochromes and demonstrating that, although endothelial cells are
capable of phagocytosis, this mechanism plays only a marginal role
in this model (Figure 3 and supplemental Figures 3-4). We
observed, as previously described, important release of unbound
CFSE within the first hours after staining31 (data not shown). Of
interest, this phenomenon did not lead to significant transfer, at
least in vitro. Conversely, no such fluorescence intensity drop was
observed for lipophilic dyes. Considering the nature of lipophilic
dyes, which become fluorescent in a lipidic environment, and
knowing that exchanges of microdomains of cell membrane do
occur, we believe it is an effective mechanism for lipophilic dye
transfer. Direct contacts between donor and acceptor populations
are not necessary for the transfer to occur, reinforcing this
hypothesis (Figure 5). Altogether, our data suggest that lipophilic
dyes induced MEC occurring through exchanges of membrane
microdomains via cell-cell interaction, a mechanism described in
the immune system as trogocytosis,34 and at distance (without
cell-cell contacts) probably involving microvesicle and/or exosome35,36 transfer. Of interest, recent reports demonstrated that
within an in vitro reconstructed model of human osteoblastic stem
cell niche, exchanges of membranes could be a new way of
reciprocal communication between HSCs and their niche.37 This
suggests that potentially any type of membrane-based labeling
strategy could possibly give rise to MEC, a hypothesis that will
need to be assessed in future studies.
In recent developments of in vivo imaging, DiI, DiD, or DiR
lipophilic dyes have been widely used for cell tracking.19-23,27,32,33 This
strategy is currently recognized as a validated—not to mention, state of
the art—technique for in vivo tracking of individual hematopoietic stem
cells.28,39 Although careful validation of biocompatibility was performed,27 our work is the first to demonstrate the importance of
optimizing the staining procedure for each cell type and, more importantly, to verify whether the fluorescent signal actually remains associated with the cells of interest in vivo and in vitro. Although the duration
of incubation before imaging is a key factor to limit dye transfer, our in
vitro data showed that it can be massive during short overnight
incubations, thus pleading for the systematic implementation of rigorous
controls to document and quantify the occurrence of MEC. Careful
reexamination of the published data using lipophilic dye–based cell
tracking allowed us to identify at least 3 situations where MEC could be
involved: long-term tracking, analysis of cell division kinetics, and
phagocytosis. Although this does not impede on the general message of
this article, it is likely that MEC did occur 70 days after the transplantation of HSPCs stained with 10␮M DiR for 30 minutes, even though it
was performed in complete medium, which reduces the staining
efficiency (data not shown).19 More recently, we noticed small discrepancies among 3 studies concerning the kinetics of cell divisions in the
first hours after injection in irradiated recipients. Indeed, 2 of these
PITFALLS ASSOCIATED WITH LIPOPHILIC DYES TRANSFER
5353
studies report that only a minority of eGFP or CFSE-labeled HSPCs
actually divide within the first 2 days after injection in conditioned
recipients.40,41 Surprisingly, a study using DiD and DiI (5␮M and
7.5␮M, respectively, for 10 minutes at 37°C in PBS without serum) to
track highly purified stem cells in vivo reports that from day 1 onward,
the majority of clusters are constituted of 2 or more cells, while nicely
demonstrating in parallel that HSCs always seeded a given BM place
individually. At the present time, it is difficult to determine whether this
discrepancy among the studies is related mainly to different kinetics of
divisions due to functional differences between the calvarium versus
long bones niches and/or whether an apparent multiplication of cells
could arise from MEC phenomenon.27 Another very impressive tracking
of human HSPCs mostly realized in nonirradiated SCID mice looked at
the impact of leukemia development on the localization of normal
human HSPCs. Considering that SCID mice maintain a relatively high
degree of immune competence (especially in the phagocytic compartment), and based on recent confirmations concerning the critical
necessity of preconditioning for engraftment to occur27,40,42 and personal
observations (F.L., manuscript in preparation), it would not be surprising
if a relatively high level of leakage (through phagocytosis and/or MEC)
occurred from human cells stained with 12.5␮M DiR for 30 minutes at
37°C. This highlights the need for further validation of some of these
observations (specially those obtained at later time points), which might
concern endogenous phagocytes rather than human HSPCs.32
During the course of this study, we have established that both
optimization of the labeling procedures and thorough control of dye
transfer are critically needed when establishing tracking experiments
involving lipophilic dyes, both in vitro and in vivo. A variety of
parameters have to be considered: the concentration of the dye (and its
vehicle), duration, temperature, and quality of the medium used during
the staining step (compare supplemental Figures 1, 4, 5, and 6).
Obviously, the washing procedure is also critical. The viability of the
stained cells, fluorescence intensity, stability, and homogeneity of the
staining over time should also be evaluated. It is also important in the
initial phases of development to analyze whether the functionality of the
cells is affected by the staining procedure. To accurately assess dye
transfer, it is important to start the flow cytometric analysis by gating
first the whole population of fluorescent cells (positive for the dye),
without prediscrimination of the population of interest (compare results
in Figure 2). The intensity of the transfer can be precisely quantified. To
determine the contribution of MEC in any model, it would be suitable to
systematically measure the amount of fluorescence associated with the
cells of interest versus other cell types (compare Figures 2D, 3, 4C, and
supplemental Figures 4, 5C). However, such quantification requires
obtaining a clearly identifiable population by flow cytometry, a condition that might not be fulfilled when small numbers of highly purified
stem cells are injected systemically.27,28,33
Thus, the main conclusion here is not only to inform the community
about the risk of dye contamination and MEC, which might have led to
misinterpretation of some results, but also to propose a simple, reliable,
and quantitative approach for systematically assessing the specificity of
any staining procedure that might be useful in the development of
improved cell-tracking models, a critical step for the quality management of cell therapy imaging.29,30,39,43-47
Acknowledgments
The authors are grateful to Dr Shai Senderovich and Christopher
Ridler, and the Animal Care Facility, the Flow Cytometry, the
Equipment Park, and the Light Microscopy core facilities at
London Research Institute for valuable technical help. We acknowledge Dr Erik Sahai for providing pivotal technical help.
From www.bloodjournal.org by guest on December 22, 2014. For personal use only.
5354
BLOOD, 1 JULY 2010 䡠 VOLUME 115, NUMBER 26
LASSAILLY et al
This work was funded by CRUK and by a European grant
(contract no. 037632; D.B.).
Authorship
Contribution: F.L. led the conception and design of the experiments, collection, analysis, and interpretation of the data, and wrote
the paper; E.G. provided material for the study, and helped in
conception and design of experiments; and D.B. helped in conception and design, data analysis, interpretation, and redaction of
paper.
Conflict-of-interest disclosure: The authors declare no competing financial interests.
Correspondence: Dominique Bonnet, Hematopoietic Stem Cell
Laboratory, CRUK, London Research Institute, 44 Lincoln’s
Inn Fields, London, WC2A 3PX; United Kingdom; e-mail:
[email protected].
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From www.bloodjournal.org by guest on December 22, 2014. For personal use only.
2010 115: 5347-5354
doi:10.1182/blood-2009-05-224030 originally published
online March 9, 2010
''Microenvironmental contaminations'' induced by fluorescent lipophilic
dyes used for noninvasive in vitro and in vivo cell tracking
Francois Lassailly, Emmanuel Griessinger and Dominique Bonnet
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