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Experimental and Molecular Pathology 96 (2014) 367–374
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Experimental and Molecular Pathology
journal homepage: www.elsevier.com/locate/yexmp
Comparative analysis of innate immune system function in metastatic
breast, colorectal, and prostate cancer patients with circulating
tumor cells
Mark F. Santos a,⁎, Venkat K.R. Mannam a, Barbara S. Craft b, Louis V. Puneky b, Natale T. Sheehan b,
Robert E. Lewis a, Julius M. Cruse a
a
b
Department of Pathology, University of Mississippi Medical Center, Jackson, MS 39216, USA
Division of Oncology, Department of Medicine, University of Mississippi Medical Center, Jackson, MS 39216, USA
a r t i c l e
i n f o
Article history:
Received 31 March 2014
and in revised form 4 April 2014
Available online 13 April 2014
Keywords:
Innate immunity
Circulating tumor cells
Toll-like receptors
Natural killer cells
Metastatic cancer
a b s t r a c t
In recent years, circulating tumor cells (CTCs) in metastatic cancer patients have been found to be a promising
biomarker to predict overall survival and tumor progression in these patients. A relatively high number of
CTCs has been correlated with disease progression and poorer prognosis. This study was designed to assess innate immune system function, known to be responsible for the immune defense against developing neoplasms,
in metastatic cancer patients with CTCs. Our aim is to provide a link between indication of poorer prognosis,
represented by the number of CTCs to the cytotoxic activity of natural killer cells, an important component
of the innate immune system, and to represent a promising expanded approach to management of metastatic
cancer patients with CTCs. Seventy-four patients, with metastatic breast, colorectal, or prostate cancer, were recruited for this study. Using a flow cytometric assay, we measured natural killer (NK) cell cytotoxicity against
K562 target cells; and CTCs were enumerated using the CellSearch System. Toll-like receptors 2 and 4 expression
was also determined by flow cytometry.
We found that within each of our three metastatic cancer patient groups, NK cell cytotoxic activity was decreased
in patients with a relatively high number of CTCs in peripheral blood compared to patients with a relatively low
number of CTCs. In the breast and prostate cancer group, patients with CTCs greater than 5 had decreased NK cell
cytotoxicity when compared to patients with less than 5 CTCs. In the colorectal cancer group, we found that 3 or
more CTCs in the blood was the level at which NK cell cytotoxicity is diminished.
Additionally, we found that the toll-like receptors 2 and 4 expression was decreased in intensity in all the metastatic cancer patients when compared to the healthy controls. Furthermore, within each cancer group, the
expression of both toll-like receptors was decreased in the patients with relatively high number of CTCs, i.e. greater
than 5 for the breast and prostate cancer group and greater than 3 for the colorectal cancer group, compared to the
patients with relatively low number, i.e. less than 5 or 3, respectively. Treatment options to increase NK cell cytotoxic activity should be considered in patients with relatively high numbers of CTCs.
Published by Elsevier Inc.
Introduction
The present investigation was designed to evaluate innate immune
system function, particularly the cytotoxicity of natural killer (NK)
cells against tumor cells. Metastatic breast, prostate, and colorectal
cancer patients were recruited for the study and each was evaluated
based on the number of circulating tumor cells (CTCs) to provide a
link between indications of poorer prognosis to the cytotoxic activity
of NK cells. CTCs are believed to be indicators of residual disease and
to portend an increased risk of metastasis and poorer outcomes for
those patients with CTCs (Plaks et al., 2013). Since the major cause
⁎ Corresponding author.
E-mail address: [email protected] (M.F. Santos).
http://dx.doi.org/10.1016/j.yexmp.2014.04.001
0014-4800/Published by Elsevier Inc.
of cancer-associated mortality is tumor metastasis, it is important to
monitor the numbers of CTCs in metastatic cancer patients. CTCs are
formed from tumor cells that separated from the primary tumor and
intravasate into blood vessels or lymphatics. These cells are then free
to migrate to different sites in the body, extravasate out of the blood
vessels, and adapt to the new microenvironment. Eventually, they will
seed, proliferate, and colonize to form metastases.
According to the National Cancer Institute, based on recent data, the
number of new cases of colon and rectum cancer was 45.0 per 100,000
men and women per year, with the number of deaths at 16.4 per
100,000 men and women per year. For breast cancer, the number of
new cases was 123.8 per 100,000 women per year and the number of
deaths was 22.6 per 100,000 men and women per year. Lastly, the number of new cases of prostate cancer was 152.0 per 100,000 men per year.
368
M.F. Santos et al. / Experimental and Molecular Pathology 96 (2014) 367–374
The number of deaths was 23.0 per 100,000 men and women per year.
These present a multitude of medical and financial issues. With such
large numbers, it is important to elucidate more effective ways to treat
cancer.
This study focuses on the function of the innate immune system,
which is believed to be the primary response against tumor growth
and development. Natural killer (NK) cells are an important component
of this system and play a vital role in eliminating cancer cells. NK cells
recognize any cell lacking the Major Histocompatibility Complex
(MHC) class I marker and release proteins that mediate the killing of
the targeted cells (Bellora et al., 2013). When functioning properly, NK
cells protect the host against a vast repertoire of potentially harmful
agents, including cells having undergone cellular transformation in the
process of forming a tumor. Other components of the innate immune
system are the Toll-like receptors (TLRs), which are found in a number
of cells including NK cells. TLRs recognize antigens that are not normally
expressed within the host, including transformed cells (i.e. cancer cells).
Upon recognition, TLRs promote the release of cytokines, chemokines,
and other agents required in antitumor immunity (Erridge, 2010).
In this study, TLRs were characterized on immune cell subsets using
monoclonal antibodies by flow cytometric methods. In addition, patient
lymphocytes were isolated from the peripheral blood and NK cell cytotoxic activity was measured using the flow cytometry assay (Bottley
et al., 2007; Bryceson et al., 2010). This study aims to assess the effectiveness of the immune system against cancer in the presence of CTCs in the
blood and to represent a promising expanded approach to management
of metastatic cancer patients with CTCs.
Materials and methods
Subjects
Seventy-four patients visiting the Multispecialty Care Cancer Clinic
at the Jackson Medical Mall in Jackson, MS were recruited for this
study. Each patient had been previously diagnosed with metastatic
breast, colorectal, or prostate cancer. A co-investigator from the study
discussed the risks and benefits, privacy issues, and other pertinent
information with potential participants before obtaining informed
consent. After obtaining consent, peripheral blood was drawn via venipuncture into a 6.0 mL sodium-heparin tube for flow cytometry analysis
and a 10 mL CellSave tube (purchased from Janssen Diagnostics, LLC)
for circulating tumor cell enumeration. At least four milliliters and
nine milliliters of blood samples are needed for the flow cytometry
analysis and CTC enumeration, respectively. In addition, a total of ten
healthy control were recruited for this study. Blood sample analysis
was also conducted using flow cytometry.
Circulating tumor cell enumeration
The CellSearch System (Janssen Diagnostics, LLC) was used to identify the number of CTCs per 7.5 mL of peripheral blood collected in
CellSave tubes. A 5 or greater CTCs per 7.5 mL whole blood is considered
the threshold at which long term prognosis and survival of breast and
prostate cancer patients is significantly diminished (Cristofanilli et al.,
2005; Hayes et al., 2006; De Bono et al., 2008). In comparison, a 3
or greater CTCs per 7.5 mL whole blood is the threshold for colorectal
cancer (Cohen et al., 2008). The CellSearch CTC test and control kits
contain immunofluorescent reagents to capture and identify CTCs of
epithelial origins. Before adding reagents, the CellSave tubes were
inverted at least 5 times to ensure thorough mixing of the preservatives
contained in the tubes with the blood samples. 7.5 mL of blood are taken
and transferred into a 15-mL conical tube included in the test kit. 6.5 mL
of dilution buffer is added and the tubes were capped and inverted at
least 5 times. The tubes were then centrifuged for 10 min at 1800 rpm
with no brake. The control was prepared following the manufacturer's
guidelines. After centrifugation, the samples were loaded onto the
CELLTRACKS® AUTOPREP® System as well as the reagents in the kit.
Ferrofluid consisting of nanoparticles with a magnetic core captures
cells expressing the epithelial cell adhesion molecule (EpCAM) for further examination. After immunomagnetic capture and enrichment,
fluorescent staining reagents were added to visualize CTCs. Fluorescent
staining reagents of anti-cytokeratin (CK) conjugated to PE, 4′-6Diamidino-2-phenylindole (DAPI), and anti-CD45 conjugated with
APC were used to label the intracellular protein cytokeratin (specific
for epithelial cells), cell nucleus, and leukocytes, respectively (all reagents were obtained from Janssen Diagnostics, LLC). After processing,
the samples were loaded onto the CELLTRACKS ANALYZER II® for
analysis of the stained cells. Positive CTCs expressed EpCAM, CK, and
DAPI and did not express CD45.
Peripheral blood lymphocyte isolation and activation
Peripheral blood lymphocytes (PBLs) were isolated from whole
blood samples using Ficoll-Hypaque centrifugation. After inverting the
sodium-heparin tube gently for 2–3 times to ensure thorough mixing
of anticoagulant to whole blood, blood was centrifuged for 5 minutes
at 3000 rpm. The resulting buffy coat was extracted using a transfer pipette and placed on a fresh tube. Approximately 2–5 mL PBS (purchased
from Sigma-Aldrich) was added to the buffy coat and mixed well. A
9 inch pipette was placed on the tube. Using a syringe, 5 mL of Ficoll
(purchased from GE Healthcare) were measured and added into the
tube through the 9 inch pipette. This resulted in the Ficoll settling to
the bottom of the tube. The tube was then centrifuged for 25 minutes
at 2000 rpm with no brake at room temperature. The resulting mononuclear cell layer, interfaced between the two layers of blood plasma and
Ficoll reagent, was removed with a transfer pipette and placed into a
fresh, clean tube. PBS was added to wash and remove remaining traces
of the Ficoll reagent. The tube was then centrifuged for 6 minutes
at 1800 rpm. The supernatant was discarded and the cell button was
resuspended in RPMI 1640 supplemented with 10% FCS (purchased
from Gibco). Cells were concentrated at 106 cells/mL and distributed
into 2 wells of a 12-well round bottom plate. Both wells contained
100 μL of 10 μg/mL LPS (purchased from Invivogen) to activate the
toll-like receptors 2 and 4. The cells were then incubated at 37 °C in
5% CO2 for 16 hours.
Immunofluorescent staining of peripheral blood lymphocytes
After the 16-hour incubation with LPS, the PBLs were prepared for
flow cytometry by immunofluorescent staining of selected CD surface
antigens and toll-like receptors 2 and 4. Two clean tubes were labeled
1 and 2, and to each, the incubated PBLs were transferred. Tube 1 was
for analysis of toll-like receptor expression. Tube 2 was used for coculturing with the target cells (K562) to measure the cytotoxic activity
of the effector cells (NK cells). Antibodies specific to selected CD markers
conjugated to different fluorochromes were purchased from Beckman
Coulter. Antibodies directed against the toll-like receptors 2 and 4
were obtained from Biolegend.
To tube 1, antibodies directed against the CD surface antigens and
the toll-like receptors were added in the following amount: 10 μL of
CD45 conjugated with AlexaFluor 700, 10 μL of CD56 conjugated with
APC, 10 μL of CD3 conjugated with ECD, 10 μL of CD14 conjugated
with PC5.5, 5 μL of anti-human TLR2 antibody conjugated with FITC,
and 5 μL of anti-human TLR4 antibody conjugated with PE. To tube 2,
the following antibodies were added: 10 μL of CD45 conjugated with
AlexaFluor 700, 10 μL of CD56 conjugated with APC, and 20 μL of CD3
conjugated with PE. Tubes were briefly vortexed and incubated in the
dark for 30 minutes at room temperature.
After incubation, 2 mL of PBS were added to each tube. The tubes
were then centrifuged for 5 minutes at 1800 rpm and the supernatant
discarded. PBS was again added and the process was repeated twice
more. After the final wash, 1.5 mL of PBS was added to tube 1. The
M.F. Santos et al. / Experimental and Molecular Pathology 96 (2014) 367–374
369
O1
O2
O3
O4
130
10 3
CD56 APC
10 2
10 1
10
0
10
0
10
1
10
2
10
3
TLR2 FITC
10 0
101
10 2
10
3
CD3 ECD
Fig. 1. Flow cytometry plot showing cell percentages of CD56+ NK cell versus CD3+ T
cells in the lymphocyte population of a metastatic cancer patient.
tube was then vortexed gently and loaded onto the flow cytometer. For
tube 2 after the final wash, the cell button was resuspended in RPMI
1640 supplemented with 10% FCS. The cells were concentrated at 106
cells/mL and co-cultured with K562 at 37 °C in 5% CO2 for 6 hours.
Co-culture of peripheral blood lymphocytes with the target cell
An effector-to-target (E:T) ratio of 20:1 was used to measure the cytotoxic activity of NK cells. Prior to co-culture, K562 cells (obtained from
the American Type Culture Collection) were immunofluorescently
stained with antibodies directed against the CD markers specific to
K562. K562 was transferred into 2 tubes (labeled tube 3 and tube 4)
and each was washed twice with PBS. Tube 3 was for measuring the
spontaneous lysis of K562 without the effector cells, while tube 4 was
used for the co-culture assay. To each tube, the following antibodies
were added: 10 μL of CD45 conjugated with AlexaFluor 700, 10 μL of
CD71 conjugated with A750, and 20 μL of CD33 conjugated with FITC.
The tubes were then incubated in the dark at room temperature for
30 minutes. After incubation, the cells were washed with PBS similar
to that mentioned above. After the final wash, 50 μL of PBS and 20 μL
of 7AAD were added to tube 3. The tube was vortexed gently and incubated for 10 minutes in the dark. Afterwards, an additional 1.5 mL
of PBS was added and the tube was vortexed gently and loaded onto
3
R1
R2
R3
R4
the flow cytometer. For tube 4 after the final wash, the cells were resuspended in RPMI 1640 supplemented with 10% FCS. They were then concentrated at 5 × 104 cells/mL to achieve the 20:1 E:T ratio and cocultured with the cells from tube 2 (mentioned above). Staining prior
to co-culture was necessary to differentiate NK cells from K562 cells
when analyzed by flow cytometry. After the 6-hour incubation, the
cells were transferred into a fresh, clean tube and washed twice with
PBS. After the final wash, 50 μL of PBS and 20 μL of 7AAD (purchased
from Beckman Coulter) were added. The tube was vortexed gently
and incubated for 10 minutes in the dark. Next, an additional 1.5 mL of
PBS was added and the tube was vortexed gently and loaded onto the
flow cytometer.
Flow cytometric detection and analysis
Cells were analyzed using Beckman Coulter flow cytometer. Positive
expression for samples containing fluorescent-labeled antibodies to CD
markers and TLRs was indicated by cells grouping into the appropriate
1st, 2nd, or 4th quadrants of their respective dot plots. The percent
positivity of cell markers was recorded in relation to the entire population (Figs. 1 and 2).
To measure the intensity of TLR expression (tube 1), cells were gated
for monocytes by plotting CD45 expression on the x-axis and CD14 +
monocytes expression on the y-axis. TLR expression was then measured
by plotting a separate histogram for the gated monocytes (Figs. 3, 4, 5,
and 6). The x-mean of the curve was taken to represent the intensity
of TLR expression.
150
10
Fig. 3. Histogram depicting TLR2 expression in the monocyte population of metastatic
cancer patients. Intensity is measured by calculating the x-mean.
CD56 APC
102
101
100
100
101
10
2
10
3
CD3 ECD
Fig. 2. Flow cytometry plot showing cell percentages of CD56+ NK cell versus CD3+ T
cells in the lymphocyte population of a healthy control sample.
10
0
10
1
10
2
10
3
TLR2 FITC
Fig. 4. Histogram depicting TLR2 expression in the monocyte population of healthy control
group. Intensity is measured by calculating the x-mean.
M.F. Santos et al. / Experimental and Molecular Pathology 96 (2014) 367–374
Gate 1
FORWARD SCATTER
30
1023
370
0
101
TLR4 PE
10
2
10
3
0
10
0
1023
SIDE SCATTER
Fig. 5. Histogram depicting TLR4 expression in the monocyte population of metastatic
cancer patients. Intensity is measured by calculating the x-mean.
Fig. 7. Flow cytometry plot of NK cells co-cultured with the target cells. Gate 1 is placed to
exclude cells with high forward and side scatter characteristics, indicative of cell debris.
To measure the NK cytotoxicity (tube 2), the percentage of target
cells killed by the effector cells was measured by the fluorescence of
7AAD in cells belonging to the CD71 labeled population. A forward
versus side scatter was first plotted (Fig. 7). Gate 1 was then placed to
exclude cells with very high forward and side scatter characteristics,
indicative of cellular aggregates or debris. Next, a dot plot of CD71
versus CD3 was generated (Fig. 8). CD3 was used as a negative selection
for K562, which were CD71 positive (Hoshino et al., 1991). Gate 2 was
then placed on the cells positive only for CD71 and a histogram was
generated based on the gated CD71 population (Fig. 9). The percentage
of dead cells was then determined by placing a marker above the 7AAD
positive cells. The fluorescence of 7AAD was also measured in the sample containing only K562 target cells (tube 3) to evaluate the percentage
of cells that lysed spontaneously. The percentage of NK cytotoxic
activity was calculated as: % cytotoxicity = ((% dead target cells in the
sample containing both effector and target cells − % spontaneously
dead target cells)/(100 − % spontaneously dead target cells)) × 100
(Kim et al., 2007).
Statistical analysis
positive. An independent t-test was conducted for each type of cancer
based on CTC profile. An ANOVA with Bonferroni multiple comparison
test was performed for all 3 types of cancer. The level of significance
was set at p b 0.05.
Results
Study population
Of the seventy-four metastatic cancer patients recruited for the
study, 36 were diagnosed with breast cancer, 30 with colorectal cancer,
and 8 with prostate cancer (Table 1). Patients with each type of neoplasm were further subdivided based on the number of CTCs present
in the blood. Patients were randomly selected without regard to sex,
age, or site of metastasis. The names of the patients were not used to
identify them. Instead, blood samples taken from patients were coded
and assigned a number according to the diagnosis: BC for breast cancer,
CR for colorectal cancer, and PC for prostate cancer.
Circulating tumor cell enumeration
Using the CellSearch System, the number of CTCs for each sample
was determined. For the breast cancer group, 25 samples had CTCs
less than 5 and 11 samples had greater than 5. For the colorectal cancer
group, 23 samples had CTCs less than 3 and 7 samples had greater than
103
150
The STATA statistics software (StataCorp) was used to analyze the
data. Samples were grouped according to the diagnosis of the patient
the blood was taken from. These were metastatic breast, colorectal,
and prostate cancer. Each group were further subdivided based on the
CTC numbers. For breast and prostate cancer, CTCs less than 5 was
taken as negative while CTCs greater 5 than was positive. For colorectal
cancer, CTCs less than 3 was negative and CTCs greater than 3 was
B1
B2
B3
B4
CD3 PE
102
101
100
10
0
10
1
10
2
10
3
10 0
Gate 2
10
1
10
2
10
3
CD71 A750
TLR4 PE
Fig. 6. Histogram depicting TLR4 expression in the monocyte population of healthy control
group. Intensity is measured by calculating the x-mean.
Fig. 8. Flow cytometry plot of CD3+ cells versus CD71+ target cells from Gate 1. Cells
positive for CD71 are shown on quadrant B4. Gate 2 is placed to select the target cell
population.
M.F. Santos et al. / Experimental and Molecular Pathology 96 (2014) 367–374
371
30
20
18
16
14
12
Dead Cells
10
8
6
4
10
0
10
1
10
2
10
2
3
7AAD
0
control
3. Lastly, for the prostate cancer group, 5 samples had CTCs less than
5 and 3 samples had greater than 5 (Table 1).
Peripheral blood lymphocyte analysis by flow cytometry
Cells were identified based on the expression of the CD antigens specific to the type of cell. Lymphocytes were gated as CD45+ and either
CD3+, for the T cell subsets, or CD56+, for the NK cell subsets. Monocytes were gated as CD45 + and CD14 +. The number of NK cells
(presented as percentage of NK cells in the lymphocyte population)
was increased in all 3 groups of cancer patients compared to the healthy
control group (Fig. 10). NK cells in the breast cancer group were found
to be significantly increased (15.90 ± 3.14, n = 36, p b 0.05) compared
to the control group (9.44 ± 1.95, n = 10). Similarly, in the colorectal
cancer group, NK cells were increased significantly (15.20 ± 3.58,
n = 30, p b 0.05) compared to the control group (9.44 ± 1.95, n = 10).
And lastly, in the prostate cancer group, NK cells were also increased
significantly (14.72 ± 3.22, n = 8, p b 0.05) compared to the control
(9.44 ± 1.95, n = 10).
Additionally, the percentage of NK cells was analyzed within
each type of cancer based on the number of CTCs (Fig. 11). For each
type of cancer, no statistical difference was found between patients
with CTCs N 5 (3 for colorectal) and patients with CTCs b 5 (3 for
colorectal). However, ANOVA test comparison of all three types of
cancer showed that there was a statistically significant difference
(n = 74, p b 0.05) in patients with CTCs N 5 (or 3) compared to patients
with CTCs b 5 (or 3).
Table 1
Study population.
Breast cancer
Colorectal cancer
Prostate cancer
Control
CTC b 5 or 3
CTC N 5 or 3
36
30
8
10
25
23
5
11
7
3
prostate
expression were also decreased significantly compared to the control
group TLR2 (10.95 ± 1.24, n = 10) and TLR4 (2.89 ± 1.15, n = 10).
Lastly, in the prostate cancer group, the intensity of TLR2 expression
(9.18 ± 1.01, n = 8, p b 0.05) was decreased significantly compared
to the control group TLR2 (10.95 ± 1.24, n = 10); however, TLR4 expression (1.92 ± 0.83, n = 8, p N 0.05) was only marginally decreased
compared to the TLR4 in the control group (2.89 ± 1.15, n = 10).
Within each type of cancer, TLR2 and TLR4 expression was analyzed
based on the number of CTCs (Figs. 13 and 14). In the breast cancer
group, the intensity of TLR2 expression was decreased significantly in
patients with CTCs N 5 (8.09 ± 2.04, n = 11, p b 0.05) compared to patients with CTCs b 5 (9.55 ± 1.88, n = 25). However, no statistical difference was observed in the expression of TLR4 (CTCs N 5: 1.26 ± 0.11,
n = 11; CTCs b 5: 1.71 ± 0.73, n = 25; p N 0.05). In the colorectal
cancer group, there was no statistical difference in the expression
of both TLR2 and TLR4 in patients with CTCs N 3 (TLR2: 7.94 ± 1.77,
n = 7, p N 0.05; TLR4: 1.21 ± 0.10, n = 7, p N 0.05) compared to
patients with CTCs b 3 (TLR2: 9.20 ± 1.59, n = 23; TLR4: 1.74 ± 0.72,
n = 23). Also in the prostate cancer group, no statistical difference
was found between patients with CTCs N 5 (TLR2: 8.34 ± 0.69, n = 3,
p N 0.05; TLR4: 1.28 ± 0.09, n = 3, p N 0.05) and patients with
CTCs b 5 (TLR2: 9.68 ± 0.83, n = 5; TLR4: 2.30 ± 0.84, n = 5).
Differences in the intensity of expression of both TLR2 and TLR4
among all 3 cancers were also analyzed using ANOVA test. TLR2 expression, for all three types of cancer, was found to be statistically different
in patients with CTCs N 5 (or 3) compared to patients with CTCs b 5
Mean NK Percentage
Breast
N
colorectal
Fig. 10. The mean percentage of natural killer (NK) cells in the control, breast cancer,
colorectal cancer, and prostate cancer group.
Analysis of TLR2 and TLR4 expression intensity by flow cytometry
TLR2 and TLR4 expression on CD45 +/CD14 + monocytes in the
cancer patients were decreased in intensity compared to the healthy
control group (Fig. 12). In the breast cancer group, the intensity
of both TLR2 (9.10 ± 2.02, n = 36, p b 0.05) and TLR4 expression
(1.58 ± 0.65, n = 36, p b 0.05) were decreased significantly compared to the control group TLR2 (10.95 ± 1.24, n = 10) and TLR4
(2.89 ± 1.15, n = 10). In the colorectal cancer group, the intensity of
TLR2 (8.91 ± 1.70, n = 30, p b 0.05) and TLR4 (1.62 ± 0.67, n = 10)
breast
0 2 4 6 8 10 12 14 16 18 20 22
Fig. 9. Histogram of CD71+ target cells from Gate 2 depicting fluorescent of 7AAD. The
percentage of dead cells is taken by placing a gate above the 7AAD peak.
Colorectal
Prostate
Type of Cancer
CTC < 5 or 3
CTC > 5 or 3
Fig. 11. Comparison of NK cell percentages in the metastatic breast, colorectal and prostate
cancer patients.
4
2
0
control
colorectal
breast
TLR2
3
2.5
2
1.5
6
1
8
.5
10
0
Mean Intensity of TLR4 Expression
12
3.5
M.F. Santos et al. / Experimental and Molecular Pathology 96 (2014) 367–374
Mean Intensity of TLR Expression
372
Breast
Colorectal
TLR4
CTC < 5 or 3
Fig. 12. Mean intensity of toll-like receptors 2 and 4 expression on the monocyte population in the control, breast cancer, colorectal cancer, and prostate cancer group.
Prostate
Type of Cancer
prostate
CTC > 5 or 3
Fig. 14. Comparison of TLR4 expression intensity in the metastatic breast, colorectal, and
prostate cancer patients.
(or 3) (p b 0.05). However, when comparing each type of cancer, no statistical difference was found (p N 0.05). In comparison, TLR4 expression
yielded similar results. That is, there was a statistically significant difference in patients with CTCs N 5 (or 3) compared to patients with CTCs b 5
(or 3) (p b 0.05) but no significant difference between each type of cancer (p N 0.05).
comparing NK cell cytotoxic activity among the individual types of
metastatic cancer, however, no statistically significant difference was
observed (p N 0.05).
Analysis of NK cell cytotoxic activity by flow cytometry
Circulating tumor cells represent a poorer prognosis and an increased risk of further metastases in previously diagnosed patients.
The complexity of cancer alone results in the ever increasing number
of studies in understanding the inner workings of the microenvironment of a tumor. CTCs are just another factor in the development
of metastatic cancer. Recent studies in breast, colorectal, and prostate
cancer have shown that in metastatic patients with a relatively high
numbers of CTCs, clinical features of cancer become more pronounced
over time, requiring patients to undergo more aggressive treatments
(Cohen et al., 2008; Cristofanilli et al., 2005; De Bono et al., 2008;
Hayes et al., 2006). The enumeration of CTCs has also been shown
to be clinically significant in monitoring patient progression in other
types of cancer, including ovarian, testicular, lung, liver, pancreatic,
head and neck, bladder cancer and melanoma (Jin et al., 2013;
Lianidou et al., 2014; Nastaly et al., 2014). Furthermore, in a previous
study of breast cancer patients, it was found that the immune function
of the NK cells was decreased significantly (Green et al., 2013). This
necessitates further elucidation of the function of the immune cells in
the tumor microenvironment in patients with high numbers of CTCs.
Breast
Colorectal
Prostate
Type of Cancer
CTC < 5 or 3
CTC > 5 or 3
Fig. 13. Comparison of TLR2 expression intensity in the metastatic breast, colorectal, and
prostate cancer patients.
14
12
10
8
6
4
2
0
Mean % Lysis of NK Cell Cytotoxicity
12
10
8
6
4
2
0
Mean Intensity of TLR2 Expression
NK cell cytotoxic activity was measured by the fluorescence of the
7AAD viability dye. Fig. 15 shows the comparison among the 3 types
of cancer by CTCs. In the breast cancer group, NK cell cytotoxicity in patients with CTCs N5 (8.28 ± 1.75, n = 11, p b 0.05) was decreased significantly compared to patients with CTCs b5 (9.65 ± 1.73, n = 25). In
comparison, NK cell cytotoxicity in the colorectal cancer group was
decreased significantly in patients with CTCs N 3 (8.95 ± 1.27, n = 7,
p b 0.05) compared to patients with CTCs b 3 (10.94 ± 2.36, n = 23).
And lastly, in the prostate cancer group, NK cell cytotoxicity was also
decreased significantly in patients with CTCs N 5 (7.85 ± 0.42, n = 3,
p b 0.05) compared to patients with CTCs b 5 (10.69 ± 1.81, n = 5).
Additionally, an ANOVA test comparison on all three types of cancer
was performed. Pooled data of NK cell cytotoxicity in all metastatic
patients showed that there was a statistically significant difference
(p b 0.05) between the patients with CTCs greater than the threshold
level (5 for breast and prostate cancer; 3 for colorectal cancer) compared to patients with CTCs less than the threshold level. When
Discussion
Breast
Colorectal
Prostate
Type of Cancer
CTC < 5 or 3
CTC > 5 or 3
Fig. 15. Percent specific lysis of NK cell cytotoxic activity in the metastatic breast, colorectal
and prostate cancer patients.
M.F. Santos et al. / Experimental and Molecular Pathology 96 (2014) 367–374
Activated human NK cells produce the cytokine IFN-γ and are able to
attack cancer cells, represented in this study by the lysis of the K562 cell
line (Souza-Fonseca-Guimaraes et al., 2012). Our results show that the
number of NK cells is increased in all three groups of cancer patients.
This suggests that these patients' NK cells are proliferating, and are
attempting to defend against the cancer. However, the NK cells cytotoxic activity is diminished significantly. The inability of the NK cells
to mount an effective antitumor response may lead to poorer prognosis
for these patients (Halama et al., 2011). Many factors responsible for the
reduction of NK cell cytotoxicity have been previously recognized
(Jewett and Tseng, 2011). Moreover, studies have shown that NK cells
obtained from cancer patients have significantly diminished cytotoxic
activity (Han et al., 1997; Miescher et al., 1988; Tsuboi et al., 1995).
The exact mechanism remains unclear. However, it is known that
many metastatic tumor cells constantly demonstrate a high NFκB activity (Rayet and Gélinas, 1999). In a study of oral cancer, the inhibition
of NFκB leads to an increase in NK cell cytotoxicity and secretion of
IFN-γ (Teruel et al., 2008; Tseng et al., 2010). Other factors that suppress immune cell effector function include tumor cell production of
inhibitory factors, such as IL-1β, IL-6, IL-8, IL-10, GM-CSF, VEGF, and
PGE2, and increased expression of KIR on the surface of immune cells
(Jewett and Tseng, 2011; Yang et al., 1996).
In comparing patients with a relatively high number of CTCs to patients with a relatively low number, we found that for all three types
of cancer, NK cell cytotoxic activity has diminished significantly. Considering patients with CTCs greater than 5 (for breast and prostate cancer)
or 3 (for colorectal cancer) per 7.5 mL of peripheral blood as the threshold indicating a poorer prognosis and increase risk of further metastasis,
these findings support our hypothesis that NK cell function is ineffective
in inducing the killing of tumor cells in patients with high number of
CTCs, thus jeopardizing patient prognosis. In a study with metastatic
melanoma, patients in different stages of the disease were evaluated
in terms of the NK cell cytotoxic activity (Fregni et al., 2013). It was
found that the decrease in the expression of the NKp46 receptor on
NK cells was correlated with the progression of the cancer, thus leading
to the diminished NK cell cytotoxicity. Similar results have been found
in other studies where NK cell function was decreased with increasing
disease spread (Lahat et al., 1989).
TLR, in addition to its role in bacterial and viral immunity, is known
to recognize endogenous ligands such as nucleic acids, intracellular proteins, and other cellular components (Yu et al., 2010). This recognition is
extended to aberrant cellular transformation, thus leading to antitumor
immunity (Zamai et al., 2007). In response to TLR stimuli, monocytes
act as detectors and provide activating signals for NK cells which,
by enhancing IFN-γ production, contribute to antitumor immunity
(Ehrentraut et al., 2011; Kloss et al., 2008). One study, however, showed
that TLR2 and TLR4 signaling on NK cells, but not on accessory cells such
as dendritic cells (DCs), was necessary for NK cell activation; and that
the intrinsic downstream signaling pathway was required for NK cell
activation (Martinez et al., 2010).
Monocytes are believed to have the highest expression of both TLR2
and TLR4 (Flo et al., 2001; Tadema et al., 2011). In our study, however,
we found a significant decrease in intensity of TLR2 and TLR4 expression
on monocytes in both the breast and colorectal cancer patients, even
after activation with LPS, a known potent inducer of both TLR2 and
TLR4 (Kanevskiy et al., 2013; Sabroe et al., 2002). In the prostate cancer
group, only TLR2 was decreased significantly; this may be attributable
to the relatively low number of patient samples. Our findings, therefore,
support that due to the low level expression of both of these TLRs,
recognition of tumor-specific antigens may also have been depressed
hence the impaired functionality of NK cells. Furthermore, in patients
with breast cancer and a relatively high number of CTCs, the intensity
of TLR2 expression is further reduced. This correlates with the decreased
NK cell cytotoxicity in these patients when compared to those with a
lower number of CTCs. However, regardless of the TLR2 and/or TLR4
expression in the colorectal and prostate cancer patients together with
373
CTC numbers, NK cell cytotoxicity remains weakened. Additional factors
remain to be investigated.
In conclusion, our findings show that NK cell cytotoxicity, in terms
of the number of CTCs in the blood, was diminished in patients with
a greater number than those with a lesser number among the three
types of cancer. In addition, the intensity of TLR2 and TLR4 expression
was also diminished, leading to ineffective activation of NK cells.
A high number of CTCs in metastatic patients thus poses a danger to
patient prognosis and overall survival.
Conflict of interest
The authors declare that there are no conflicts of interest.
Acknowledgements
The authors express genuine appreciation to the lab personnel
Heather Jones and Carol Irizarry-Capetillo.
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