ONCOLOGY LETTERS 22: 522, 2021

Novel circulating tumor cell‑detection chip combining
conventional podoplanin and EGFR antibodies for
all histological malignant pleural mesothelioma
MASATOSHI KANAYAMA1, RINTARO OYAMA1, MASATAKA MORI1, AKIHIRO TAIRA1, SHINJI SHINOHARA1,
TAIJI KUWATA1, MASARU TAKENAKA1, KAZUE YONEDA1, KOJI KURODA1,
TAKASHI OHNAGA2, YUKINARI KATO3,4 and FUMIHIRO TANAKA1
1

Second Department of Surgery, University of Occupational and Environmental Health, Kitakyushu,
Fukuoka 807‑8555; 2Central Research Laboratories, Toyama Industrial Technology Center, Takaoka, Toyama 933‑0981;
3
Department of Antibody Drug Development, Tohoku University Graduate School of Medicine;4
New Industry Creation Hatchery Center, Tohoku University, Sendai, Miyagi 980‑8575, Japan
Received December 7, 2020; Accepted March 18, 2021
DOI: 10.3892/ol.2021.12783
Abstract. In our previous study, a microfluidic system wasdeveloped based on podoplanin detection for capturing circu‑
lating tumor cells (CTCs), derived from malignant pleural
mesothelioma (MPM). However, non‑epithelioid MPM
shows low podoplanin protein expression compared with thatin epithelioid MPM; thus, some CTC populations may be
missed. To overcome this limitation, a new CTC‑detection
chip was developed by combining the conventional podo‑
planin antibody (clone: NZ‑1.2) with an epidermal growth
factor receptor (EGFR)‑targeted antibody (cetuximab). The
cell‑capture efficiency of the Cocktail‑chip reached 100% in
all the histological MPM cell lines. The median CTC‑counts
from 19 patients with MPM (epithelioid/non‑epithelioid: 10/9)
with the NZ‑1.2‑ and Cocktail‑chips were 1 and 3 (P= 0.311)
in 1 ml peripheral blood, 1.5 and 2 (P= 0.332) in epithelioid
MPM, and 1 and 3 (P= 0.106) in non‑epithelioid MPM, respec‑tively. Overall, the Cocktail‑chip showed an improved ability
to detect more CTCs in patients with non‑epithelioid MPM
compared with that in the conventional NZ‑1.2‑chip, showing
non‑significant, but higher CTC detection. Furthermore,
CTC‑counts, determined using the Cocktail‑chip were signifi‑
cantly correlated with the clinical stage of non‑epithelioid

Correspondence to: Dr Fumihiro Tanaka, Second Departmentof Surgery, University of Occupational and Environmental Health,1‑1 Iseigaoka, Yahatanishi‑ku, Kitakyushu, Fukuoka 807‑8555,
Japan
E‑mail: ftanaka@med.uoeh‑u.ac.jp

Abbreviations: CTCs, circulating tumor cells; EGFR, epidermal

growth factor receptor; EpCAM, epithelial cell adhesion molecule;
MPM, malignant pleural mesothelioma

Key words: circulating tumor cells, malignant pleural mesothelioma,
CTC‑chip, podoplanin, epidermal growth factor receptor

MPM. In epithelioid MPM, the Cocktail‑chip achieved a
CTC‑detection efficiency equivalent to that in the conventional
NZ‑1.2‑chip. The Cocktail‑chip enabled sensitive CTC detec‑tion of all histological MPM, including the non‑epithelioid
subtype, which may provide a foundation for the diagnosis,
treatment, and prognosis of MPM progression.
Introduction
Circulating tumor cells (CTCs) are derived from the primarytumor and circulate in the peripheral blood (1). They are consid‑
ered to play a crucial role in metastasis formation, which is theleading cause of cancer‑related death. Therefore, the detection
and molecular biological analysis of CTCs may enhance thediagnosis and treatment of patients with cancer (2‑4). Amongthe various CTC‑detection devices currently available, only
the CellSearch system (Menarini Silicon Biosystems Spa)has been approved by the Food and Drug Administration forclinical use (5), and has produced highly reproducible results
and demonstrated clinical relevance in breast, colorectal,
prostate and lung cancer (6‑11). However, as CellSearch isan epithelial cell adhesion molecule (EpCAM)‑dependentcell‑capture system, it fails to identify CTCs in non‑epithelial
tumors with low EpCAM expression, such as malignant
pleural mesothelioma (MPM) (12). To overcome this diag‑nostic gap, in our previous studies, a new microfluidic device
system, named as the ‘Universal CTC‑Chip’, was developed,
which enables EpCAM‑independent cell capture by attachingvarious antibodies to a large number of microposts on thechip surface (13,14). In addition, the clinical significance of
CTCs in MPM was found and CTC capture was possible using
an antibody against podoplanin, a well‑known diagnostic
marker of MPM (14‑16). However, podoplanin expression islower in non‑epithelioid MPM (30‑75%) compared with thatin epithelioid MPM (80‑100%) (17‑20), which may reduce
the efficiency of this diagnostic approach. Furthermore, ourprevious study showed that CTCs were detected in 92.3%(12/13 patients) of epithelioid MPM cases compared with that

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KANAYAMA et al: CTC-DETECTION CHIP FOR MALIGNANT PLEURAL MESOTHELIOMA

in 33.3% (3/9 patients) of non‑epithelioid MPM cases (16),suggesting that some CTC populations do not express podo‑
planin, particularly those of the non‑epithelioid subtype.
Epidermal growth factor receptor (EGFR) is a 170 kDa trans‑
membrane protein with intrinsic tyrosine kinase activity that
regulates cell growth (21). EGFR is overexpressed in several
malignancies, including non‑epithelioid MPM (22‑25). In
recent years, cetuximab, a chimeric antibody targeting EGFR,
was found to be an effective cell‑capture antibody comparedwith that in other EGFR antibodies, due to its low dissociation
constant and strong cell adhesion ability (26). In the present
study, the following was investigated: i) EGFR expression in
the MPM cell lines; ii) the cell‑capture efficiency of a CTC‑chipcoated with cetuximab; and iii) whether an antibody cocktail
of podoplanin (clone NZ‑1.2) and cetuximab could enhance
CTC capture in all histological subtypes of MPM.
Materials and methods
This research was conducted with the approval of the
Ethics Committee of the University of Occupational andEnvironmental Health, Japan (approval no. H26‑15).
Cell lines. A total of 5 human cell lines representing different
histological subtypes of MPM were used: Epithelioid MPM:
ACC‑MESO‑1, ACC‑MESO‑4, and NCI‑H226; biphasic
MPM: MSTO‑211H; and sarcomatoid MPM: NCI‑H28. The
ACC‑MESO‑1 and ACC‑MESO‑4 cell lines were purchasedfrom the Riken BioResource Research Center, while theNCI‑H226, MSTO‑211H, and NCI‑H28 cell lines werepurchased from the American Type Culture Collection. All thecell lines were cultured in RPMI‑1640 (FUJIFILM Wako Pure
Chemical Corporation), supplemented with 10% fetal bovine
serum (Thermo Fisher Scientific, Inc.) at 37˚C in a humidified
incubator with 5% CO2.
Flow cytometry analysis. To analyze EGFR expression, the celllines were incubated with an anti‑EGFR antibody (1:100 dilu‑tion; clone 528; cat. no. sc‑120; Santa Cruz Biotechnology,Inc.) for 1 h at room temperature. The cells were subsequently
incubated with goat anti‑mouse IgG antibody conjugated withfluorescein isothiocyanate for 30 min at room temperature(1:20 dilution; cat. no. 349031; BD Biosciences). Flow cytom‑
etry analysis was performed using an EC800 Cell Analyzer(Sony Biotechnology, Inc.), and the data was analyzed using
FlowJo software version 10 (Becton‑Dickinson and Company).
The mean fluorescence intensity was calculated as the ratio of
the positive control to that of the negative control (PBS with
1% bovine serum albumin; Nacalai Tesque, Inc.).
CTC‑chip preparation. The CTC‑chip was first coated with
antibodies in a two‑step process as previously described (14),then used for the experiments. Briefly, the CTC‑chip was
incubated with the base antibody overnight at 4˚C followed
by incubation with the capture antibody at room temperaturefor 1 h.
In the present study, the CTC‑chip was built basedon the following three combinations: i) previously
established NZ‑1.2‑chip (16) with goat anti‑rat IgG(200 µg/ml; cat. no. 3052‑01; SouthernBiotech) as thebase antibody and rat anti‑human podoplanin antibody(5,000 µg/ml; clone NZ‑1.2) (27) as the capture antibody;i) Cetuximab‑chip with goat anti‑human IgG (200 µg/ml; cat.
no. 2043‑01; SouthernBiotech) as the base antibody and cetux‑imab (5,000 µg/ml; Bristol‑Myers Squib Company) as thecapture antibody and iii) Cocktail‑chip with goat anti‑rat IgG(200 µg/ml) + goat anti‑human IgG (200 µg/ml) as base anti‑
bodies and rat anti‑human podoplanin antibody (5,000 µg/ml)+ cetuximab (2,500 µg/ml) as capture antibodies.
Sample preparation and evaluation of cell‑capture efficiency
in the MPM cell lines. Sample preparation and evaluation ofcell‑capture efficiency was performed as previously described(14). Tumor cells labeled using the CellTrace CSFE cell prolif‑
eration kit (Thermo Fisher Scientific, Inc.) were suspended in
1 ml blood collected from a healthy volunteer (obtained from
author MK; single venous blood collection from the elbow;
100 cells/ml). This sample was added to each CTC‑chipsystem at a constant flow rate (1.0 ml/h) and monitored using
a fluorescence microscope (CKX41; Olympus Corporation).
The total number of cells added to the CTC‑chip (N‑total)
was determined by counting the number of cells that passed
through the inlet of the CTC‑chip, whereas the number of
captured cells (N‑captured) was determined by counting thenumber of cells that remained on the CTC‑chip. Cell‑capture
efficiency was calculated as the N‑captured/N‑total x100 (%),
and the mean ± SE was calculated for each sample. Eachexperiment was performed in triplicate.
Clinical evaluation of CTCs in patients with MPM. Peripheral
blood samples were collected from 19 patients with MPM
between January 2018 and August 2020, to assess CTCs
at diagnosis or prior to treatment. A total of 6 ml blood wascollected in a collection tube (BD Vacutainer EDTA‑2K;
BD Biosciences), and after sufficient suspension, 1 ml blood
was added to both the NZ‑1.2‑ and Cocktail‑chips. The charac‑
teristics of the patients with MPM are summarized in Table I.
Clinical stage was determined according to the guidelines of
the International Mesothelioma Interest Group, version 8 (28).
All patients provided written informed consent to participate in
the study. The cells captured on 2 of the CTC‑chips were incu‑
bated with a primary antibody, a rabbit anti‑cytokeratin (CK)
antibody (1:100 dilution; cat. no. ab9377; Abcam) and a mouseanti‑CD45 antibody (1:100 dilution; cat. no. 304002; clone
HI30; BioLegend, Inc.) for 1 h at room temperature, followed
by incubation with 30 min of incubation at room temperature
with a secondary antibody, an Alexa Fluor 594 anti‑rabbit
IgG antibody (1:100 dilution; cat. no. A‑11037; Thermo Fisher
Scientific, Inc.) and an Alexa Fluor 488 anti‑mouse IgG antibody(1:100 dilution; cat. no. A‑11029; Thermo Fisher Scientific, Inc.)
containing 1 µg/ml Hoechst 33342 (cat. no. 4082; Cell Signaling
Technology, Inc.) Cells with round‑to‑oval morphology,
Hoechst 33342‑positive nuclei, CK‑positive staining in thecytoplasm, and CD45‑negative staining were identified as
CTCs using a fluorescence microscope at x10 magnification(DMi8‑S2G; Leica Microsystems), and CTCs were indepen‑dently identified by two investigators who were blinded to theclinical data. Survival analysis according to the CTC‑counts
in NZ‑1.2‑chip and Cocktail‑chip was also performed. The
median follow‑up was 175 (range, 28‑1,067) days.

ONCOLOGY LETTERS 22: 522, 2021

Table I. Patient characteristics (n=19).
Characteristic
Mean age (range), years
Sex, n (%)
Male
FemaleTumor laterality, n (%)
Right
Left
TNM stage, n (%)
IA
IB
II
IIIA
IIIB
IV
Histology, n (%)
Epithelioid
Non-epithelioid
Biphasic
Sarcomatous

Value
69.0 (55-78)
19 (100.0)
0 (0.0)13 (68.4)
6 (31.6)
2 (10.5)
7 (36.8)
2 (10.5)
1 (5.3)4 (21.1)
3 (15.8)
10 (52.6)
9 (47.4)
4 (44.4)
5 (55.6)

In addition, to evaluate non‑specific detection, peripheral
blood samples were collected from five healthy individuals(6 ml; single venous blood collection from the elbow) and used
to detect CTCs in the same manner as aforementioned. The
healthy volunteers were recruited from our laboratory staff
and provided formal written informed consent to participate.
Data collection from the healthy subjects was also included in
the Ethics Committee approval (approval no. H26‑15).
Evaluation of podoplanin and EGFR expression using
immunohistochemical staining in the primary lesions. Serial
4‑µm‑thick sections were cut from each 10% formalin‑fixedand paraffin‑embedded primary tumor specimen collected bypleural biopsy or radical surgery, then evaluated using immu‑
nohistochemistry staining according to standard protocols.
Sections were heated in 0.01 M citrate buffer (pH 6.0; cat.
no. RM102‑C; LSI Medience Corporation) at 98˚C for 15 minfor antigen retrieval and incubated in 3% hydrogen peroxide (cat.
no. 081‑04215; FUJIFILM Wako Pure Chemical Corporation)for 10 min to inactivate endogenous peroxidase. After blocking
with Protein Block Serum‑Free (cat. no. X090930‑2; Agilent
Technologies, Inc.) for 15 min, sections were incubated withmouse anti‑podoplanin monoclonal antibody (clone D2‑40;
cat. no. 413451; pre‑diluted antibody; Nichirei Biosciences,Inc.) and rabbit anti‑EGFR monoclonal antibody (1:50 dilu‑tion; clone D38B1; cat. no. 4267S; Cell Signaling Technology,Inc.) for 1 h. Sections were then washed and incubated with
Histofine Simple Stain MAX PO (MULTI) (cat. no. 424152;
Nichirei Biosciences, Inc.) for 30 min. Thereafter, sections
were visualized with DAB+ Liquid (cat. no. K346811; Agilent
Technologies, Inc.) for 10 min and counterstained with hema‑
toxylin for 1 min (cat. no. 30002; Muto Pure Chemicals Co.,

3Ltd.). All steps after the antigen retrieval step were performedat room temperature.
Statistical analysis. Differences between continuous variables
were evaluated using a Wilcoxon signed rank test for paired
data, that was not normally distributed. The correlation
between two variables was analyzed using Spearman's rankcorrelation analysis. The Kaplan‑Meier method was used toestimate the probability of survival, with survival differences
being analyzed using the log‑rank test. P<0.05 was considered
to indicate a statistically significant difference. All statisticalanalyses were performed using SPSS software (version 27.0;
IBM Corp.).
Results
Cell‑capture efficiency of the three types of CTC‑chip. The
flow cytometry results of EGFR expression are shown in Fig. 1.
EGFR was detected in all the cell lines, including biphasic
and sarcomatoid MPM subtypes, which have low podoplanin
expression.
The optimal concentration of cetuximab to be used on
the newly designed CTC‑chip was determined using the
NCI‑H226 cell line. The cell‑capture efficiencies were 89.2,
89.1 and 102.5% at cetuximab concentrations of 500, 1,000
and 5,000 µg/ml, respectively (data not shown). Therefore,
5,000 µg/ml was used as the optimal cetuximab concentration
for the further experiments. The cell‑capture efficiencies of thethree types of CTC‑chips are shown in Fig. 2. The epithelioid
MPM cell lines, with high podoplanin expression, were effec‑
tively captured by the NZ‑1,2‑chip containing podoplanin, but
the cell‑capture efficiency for the non‑epithelioid cell lines was
low. The Cetuximab‑chip, containing the anti‑EGFR antibody,
had high cell‑capture efficiency in the majority of the cell lines,
except for the ACC‑MESO‑4 cell line, in which there was a
40% cell‑capture efficiency. The Cocktail‑chip, containing
antibodies targeting both podoplanin and cetuximab, reached100% of a cell-capture efficiency in all the MPM cell lines;
therefore, it was used for clinical evaluation.
Clinical evaluation of the Cocktail‑chip in patients with MPM.
Representative images of immunofluorescent staining of CTCs
captured using the Cocktail‑chip in patients with malignant
pleural mesothelioma are shown in Fig. 3. Fig. 4 shows the distri‑bution of CTC‑counts using the NZ‑1.2‑ and Cocktail‑chips
for each sample. No cells were detected in the five healthy
subjects, and no non‑specific detection was observed (Fig. 4D).
CTCs were detected in 73.7% of the samples (14/19 patients)
with both the NZ‑1.2‑ and Cocktail‑chips. Furthermore, theNZ‑1.2‑ and Cocktail‑chips detected CTCs in 90 (9/10 patients)
and 70% (7/10 patients) of samples with epithelioid MPM,
and in 55.6 (5/9 patients) and 77.7% (7/9 patients) of samples
with non‑epithelioid MPM, respectively. No clusters, such as
clusters of tumor cells or clusters of tumor cells and whiteblood cells, were observed. The median CTC‑counts using theNZ‑1.2‑ and Cocktail‑chips were 1 (range, 0‑9) and 3 (range,0‑12) in the overall population (P=0.311), 1.5 (range, 0‑9) and2 (range, 0‑12) in epithelioid MPM (P= 0.332), and 1 (range,0‑4) and 3 (range, 0‑9) in non‑epithelioid MPM (P= 0.106),
respectively. There was no significant difference between the

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KANAYAMA et al: CTC-DETECTION CHIP FOR MALIGNANT PLEURAL MESOTHELIOMA

Figure 1. Expression of EGFR and podoplanin on the surface of malignant pleural mesothelioma cell lines detected using flow cytometry. EGFR, epidermal
growth factor receptor; MFI, mean fluorescence intensity.

Figure 2. Cell‑capture efficiency of the NZ‑1.2‑, cetuximab‑ and Cocktail‑chips in malignant pleural mesothelioma cell lines.

Figure 3. Representative images of immunofluorescent staining of CTCs captured using the Cocktail‑chip in patients with malignant pleural mesothelioma.
CTCs, circulating tumor cells.

ONCOLOGY LETTERS 22: 522, 2021

5

Figure 4. Wilcoxon signed rank analysis of CTCs detected using the NZ‑1.2‑ and Cocktail‑chips according to their count. The CTCs were analyzed from 1 ml
peripheral blood collected from patients with (A) malignant pleural mesothelioma (all patients), (B) epithelioid, (C) non‑epithelioid subtype and (D) healthy
subjects. CTCs, circulating tumor cells.

two chips; however, the Cocktail‑chip detected more CTCs
in non‑epithelioid MPM, suggesting it was more effective at
detecting CTCs. In addition, the Cocktail‑chip achieved a

CTC‑detection efficiency equivalent to that of the conventional
NZ‑1.2‑chip in epithelioid MPM, that is there was no change in
the number of epithelial cells (Fig. 4B).

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KANAYAMA et al: CTC-DETECTION CHIP FOR MALIGNANT PLEURAL MESOTHELIOMA

Figure 5. Spearman’s correlation analysis of CTCs detected using the NZ‑1.2‑ and Cocktail‑chips according to their count and clinical stage. The CTCs wereanalyzed from 1 ml peripheral blood collected from patients with (A) malignant pleural mesothelioma (all patients), (B) epithelioid, and (C) non‑epithelioid
subtype. CTCs, circulating tumor cells.

The correlation coefficient between CTC‑counts andclinical stage in all patients was 0.194 (P= 0.425) with the
NZ‑1.2‑Chip and 0.413 (P= 0.079) with the Cocktail‑chip.
Based on the histology data, the correlation coefficients inepithelioid and non‑epithelioid MPM were 0.522 (P= 0.121)
and 0.199 (P=0.582) for CTC‑counts with the NZ‑1.2‑Chip and
0.108 (P= 0.766) and 0.763 (P= 0.017) with the Cocktail‑chip,
respectively (Fig. 5). These data indicated that the CTC‑counts
using the Cocktail‑chip were significantly correlated with theclinical stage of non‑epithelioid MPM.

The survival analysis is shown in Fig. 6. There was no
statistically significant difference between the presence of
CTCs and prognosis in both the NZ‑1.2‑ and Cocktail‑chips;
however, patients with CTCs detected had a poorer prog‑
nosis (P= 0.274 and P= 0.114, respectively). Furthermore, inpatients with stage I MPM, one patient without CTCs detectedsurvived without progression for >3 years following surgery,whereas 2 patients with CTCs detected showed early progres‑sion and died following treatment. More specifically, case 1(NZ‑1.2‑chip:2, Cocktail‑chip:12) died 4 months following

ONCOLOGY LETTERS 22: 522, 2021

7

Figure 6. Survival analysis of CTCs in patients with malignant pleural mesothelioma. Kaplan‑Meier survival curves and log‑rank tests were used for the
analysis in patients with no CTCs (0) and ≥1 CTCs using the (A) NZ‑1.2‑ and (B) Cocktail‑chips. CTCs, circulating tumor cells.

Figure 7. Immunohistochemical staining of the primary lesions in patients with malignant pleural mesothelioma. Staining of (A) podoplanin (clone D2‑40) and(B) epidermal growth factor receptor (clone D38B1).

chemotherapy due to tumor progression, including distantmetastasis, and case 2 (NZ‑1.2‑chip:2, Cocktail chip:4) died
4 months following surgery due to locally advanced tumor
progression (data not shown).
Evaluation of podoplanin and EGFR expression using immu‑
nohistochemical staining in primary lesions. Podoplaninstaining was strongly positive in all cases of epithelioid MPM,
whereas it was negative or weakly positive in 3 out of 5 cases ofsarcomatoid MPM. In biphasic MPM, the epithelial component
was strongly positive in all four cases, whereas the sarcoma
component was negative or weakly positive in some cases. In
contrast, there were no negative cases of EGFR, and all werestrongly positive except for 2 epithelioid and 1 biphasic MPM,
which were weakly positive (Fig. 7).
Discussion
In the present study, EGFR expression was confirmed in all
types of the MPM cancer cell lines. By combining with cetux‑imab (Cocktail‑chip), the cell‑capture efficiency was higher in
non‑epithelioid MPM‑derived CTCs compared with that in theconventional NZ‑1.2‑chip, which was designed to only detect
podoplanin.

Effective capture of rare CTCs presents technical chal‑lenges. A common strategy is EpCAM‑dependent isolation, as
is used in CellSearch, because epithelial tumor cells express
EpCAM. However, EpCAM‑dependent methods may not beapplicable to the isolation of non‑epithelial tumor cells (15),and CTCs in MPM have not been fully investigated so far. In
our previous studies, a CTC capture system for MPM usinganti‑podoplanin antibodies (clone E1) (14,15) and NZ‑1.2 (16)was developed, and demonstrated its clinical significance.
However, that system did not detect CTCs in non‑epithelioid
MPM, such as sarcomatoid and biphasic subtypes, which
are particularly aggressive (29). Therefore, the present study
focused on EGFR as it is widely expressed in several types oftumor, such as gastric, liver, colorectal and lung cancer (21).
EGFR expression was observed in non‑epithelioid MPM
cell lines with poor podoplanin expression, indicating that
EGFR represents a suitable target for CTC detection. The
Cetuximab‑chip showed high cell‑capture efficiency; however,this chip exhibited a low capture efficiency for one epithelioid
subtype cell line (ACC‑MESO‑4). Thus, to overcome thispossible detection limitation of cetuximab, a new chip was
designed by combining antibodies targeting podoplanin andcetuximab. This dual detection approach had a high capture
efficiency in all histological MPM cell lines. In a clinical setting,

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KANAYAMA et al: CTC-DETECTION CHIP FOR MALIGNANT PLEURAL MESOTHELIOMA

the Cocktail‑chip also achieved a high cell‑capture efficiency,
as well as correlation with clinical stage in non‑epithelioid
subtype MPM cases. In addition, the immunostaining results
showed that there were several cases of poor expression of
podoplanin in non‑epithelioid MPM. These results suggested
that cetuximab could be used in combination with podoplanin
to support CTC detection in the non‑epithelioid subtype.
Furthermore, the Cocktail‑chip achieved a CTC‑detection
efficiency equivalent to that of the conventional NZ‑1.2‑chipin epithelioid MPM. Therefore, the results from the present
study suggested that the Cocktail‑chip could be used for CTC
detection in all histological MPM cases.
In the present study, the prognostic analysis was not
sufficient, due to the low number of cases and the observa‑
tion period was short in some cases. There was no association
between CTC detection and prognosis; however, patients inwhich CTCs were detected there was a poorer prognosis,
which may indicate that CTC detection could be of benefit intreatment selection, such as prioritizing chemotherapy oversurgery in patients in which CTCs were detected in the earlystage. In addition, distant metastasis is a rare progression
event in MPM and mainly occurs due to local metastasis;
however, based on the results for case 2 in the present study
and a reported case in which an increased number of CTCs
during CTC monitoring contributed to detection of local
progression (16), CTC detection in MPM may indicate local
progression as well as distant metastasis. These clinical find‑ings should be verified in further studies with larger numbersof cases.
The present study has several limitations. First, only
19 patients were included in this study; therefore, the cohort is
too small to draw a definitive conclusion on the efficiency of
the Cocktail‑chip. Second, CTC detection and counting was
performed only once for each patient. Therefore, collecting
peripheral blood samples from newly diagnosed and treatedpatients with MPM is currently ongoing, which will verify the
prognosis and predictive value of the general CTC‑detection
system. Third, there may have been variability in the experi‑
ment as 1 ml blood from each patient was used for each chip,and the experimental protocol was not fully automated.
Finally, in the current study, it was not validated whether thecells captured with the CTC‑chip were true MPM cells. To
resolve these system‑related limitations, the application of afully automated system using the automated pipetting system‘EDR‑24LX’ (BIOTEC Co., Ltd.) is being evaluated anddeveloping a protocol for genetically analyzing cells capturedby the CTC‑chip using the micromanipulator ‘M‑152’(NARISHIGE Group).
In conclusion, the novel Cocktail‑chip was more effectiveat capturing CTCs from all histological MPMs, including
the non‑epithelioid subtype. Further studies may reveal theclinical value of MPM‑derived CTCs captured using the
Cocktail‑chip. Based on the results from the present study, thisCocktail‑chip system may assist in the development of novel
diagnostic, therapeutic and prognostic options for monitoring
MPM progression.
Acknowledgements
Not applicable.

Funding
This study was supported by the University of Occupational
and Environmental Health Research Grant for Promotion of
Occupational Health (grant no. 902) and Japan Agency for
Medical Research and Development (AMED) under grantnos. JP20am0401013, JP20am0101078, JP20ae0101028 andJP20bm1004001.
Availability of data and materials
The datasets used and/or analyzed during the current study areavailable from the corresponding author on reasonable request.
Authors' contributions
MK, KY, TO, YK and FT contributed to the conception anddesign of the study. AT, SS, TK, MT and KK provided thesamples and analyzed the data. MK, RO and MM performed
the cell experiments. TO performed the CTC‑Chip andprovided technical support. YK provided the capture anti‑
bodies. MK drafted the manuscript. TO and YK revisedmanuscript for important intellectual content. KY and FT
supervised the study and revised the manuscript. All authors
read and approved the final version of the manuscript.
Ethics approval and consent to participate
This research was conducted with the approval of the
Ethics Committee of the University of Occupational andEnvironmental Health, Japan (approval no. H26‑15). All
patients provided written informed consent to participate in
the study.
Patient consent for publication
Not applicable.
Competing interests
The authors declare that they have no competing interests.
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