CD13 is a candidate marker closely correlated with SP cells.
To identify specific cell-surface markers that correlate with the SP fraction, we utilized our previous data sets of SP and non-SP fraction gene expression profiles obtained using microarray analyses (8
). From a list of 268 genes upregulated in the SP cells (with a fold change > 2) (8
), we selected 56 genes that potentially encode cell-surface proteins via the UniProtKB database (
). Working from the list of 56 upregulated genes (Supplemental Table 1; supplemental material available online with this article; doi:
) and an additional 43 markers reported to be closely associated with normal stem cells and CSCs, we tested 47 commercially available antibodies (Supplemental Table 2) to identify surface markers that were enriched in the SP fraction (Figure A).
CD13 is a candidate marker of the SP fraction.
During this screening, we identified 2 candidate markers, CD13 and CD31. The expression analysis of CD13 was 1.64 ± 0.45 in the SP and 0.51 ± 0.03 in the non-SP cell fraction (P < 0.01) (Figure B). We focused on CD13 in the current study, since the expression of CD31 was abundant in the G2/M/SP fraction but was not universal in the liver cancer cell lines studied by us (HuH7, PLC/PRF/5, and Hep3B), and the statistical significance was weak (P = 0.076) (Figure B and Supplemental Figure 1, A and B).
Expression of CD13, CD133, and CD90 was assessed in hepatitis infection–negative (HuH7) and –positive (PLC/PRF/5) cell lines. The expression of CD133 was detected in HuH7 but not in PLC/PRF/5, and the expression of CD90 was detected in PLC/PRF/5 but not in HuH7. The expression of CD13 was observed in both these cell lines as well as in Hep3B (Figure C and Supplemental Figure 1A). In HuH7 in particular, the CD13+ cells typically existed in a CD133strong fraction (CD13+CD133+).
Multicolor analysis with Hoechst staining exhibited clear localization of CD13+ cells in the SP fraction of HuH7 and PLC/PRF/5, whereas the CD13–CD133+ and CD90+ fractions were localized to the G1-to-G2 fraction and not the SP fraction (Figure D). To confirm the cell-cycle status of the CD13+ cells in PLC/PRF/5, cell-cycle analysis by combined multicolor analysis and 7-amino-actinomycin D (7-AAD) DNA labeling was performed. The CD13+CD90– population was mainly in the G0/G1 phase, and the CD13+CD90+ population was clearly in the S to G2/M phase. The CD13–CD90+ cells were present in all phases of the cell cycle but were more clearly present in the G2/M and S phases when compared with the CD13+CD90– population (Supplemental Figure 1C).
In these studies, we confirmed CD13 as a universal candidate marker that correlates with the liver cancer SP fraction. There were no definitive single markers that showed a stronger correlation to the SP fraction than CD13 and, to a lesser extent, CD31.
CD13 is a marker of tumor-initiating and potentially dormant HCC cells.
Given that hematopoietic and leukemic stem cells are in the G0
phase, identification and characterization of dormant or slow-growing cancer cell populations is very important because of these populations’ relevance to chemo resistance and recurrence. Studies of CD13 expression in HuH7 and PLC/PRF/5 and their relationships with the cell-cycle phase, using the DNA-binding dye Hoechst 33342 and the RNA-binding dye pyronin Y (PY) (3
), indicated that most of the CD13+
fraction exists in the G1
phase and the CD13strong
population was clearly localized in G0
. The CD133+
population in HuH7 and the CD90+
fraction in PLC/PRF/5 were distributed in the G1
/M phases, respectively. The relationships between the SP fraction and the G0
cell-cycle phase were also confirmed, and the SP fraction was clearly localized in the G0
phase under reserpine-free (ABC transporter blocker) conditions (Figure A).
CD13 is a candidate marker of dormant to slow-growing CSCs.
To study the cell fate and dye-retaining capacity of HuH7 CD13+ cells, the cell-surface membrane was labeled with PKH26GL reagent and cell fate was traced for 238 hours. Equal numbers of cells were seeded for each population. The CD13+CD133+ fraction exhibited very slow growth compared with the CD13–CD133+ fraction, with the doubling time of the CD13+CD133+ fraction estimated at approximately 160 hours. Dye-retaining cells could be observed 238 hours after cell seeding only in the CD13+CD133+ fraction (Figure B and Supplemental Videos 1–3). The CD13–CD133– fraction exhibited cell fragmentation and apoptotic changes during cell culture. To confirm CD13 expression in association with cell growth, we performed cell proliferation assays. Data from isolated HuH7 populations showed that CD13+CD133+ cells exhibited slow cell growth compared with CD13–CD133+ cells 72 hours after seeding (Figure C). The CD13–CD133– population also grew slowly but maintained viability for a week, with difficulty, because of apoptosis.
Next, tumor-formation ability of each fraction was studied in HuH7 and PLC/PRF/5 cells. Limiting dilution analysis of HuH7 cells revealed that the CD13+CD133+ fraction formed tumors from 100 cells (2/4), the CD13–CD133+ fraction formed tumors from 1,000 cells (3/4), and the CD13–CD133– fraction formed no tumors from 5,000 cells (0/4) in NOD/SCID mice after 4 weeks of observation. In PLC/PRF/5 cells, the CD13+CD90– fraction formed tumors from 100 cells (2/4), and the CD13–CD90+ fraction formed tumors from 5,000 cells (2/2), whereas the CD13–CD90– cells formed no tumors from 5,000 cells (0/4) in NOD/SCID mice after 6 weeks of observation (Table ). To assess the tumor formation ability definitively, formed tumors were digested, and isolated CD13+CD133+ and CD13–CD133+ fractions of HuH7 and isolated CD13+CD90– and CD13–CD90+ fractions of PLC/PRF/5 were serially transplanted to secondary NOD/SCID mice. As controls, nonisolated cell fractions of HuH7 and PLC/PRF/5 were also serially transplanted. Tumor formation ability of CD13+ cells compared with that of CD13– cells in serial transplantation assay was demonstrated more clearly than that of limiting dilution assay. In HuH7, after 6 weeks of observation, the CD13+CD133+ fraction formed tumors from 500 cells (1/4), whereas the CD13–CD133+ fraction and control formed very small tumors only in 10,000 cells (1/4 in the CD13–CD133+ fraction, and 2/6 in control). In PLC/PRF/5, after 6 weeks of observation, the CD13+CD90– formed tumors from 500 cells (2/4), and the control formed tumors from 5,000 cells (1/6), whereas the CD13–CD90+ fraction formed 1 very small tumor only in 10,000 cells (1/4) (Table ).
Limiting dilution and serial transplantation assay of HuH7 and PLC/PRF/5 cells
To assess semiquiescent status of CD13+ cells in vivo, BrdU-retaining status was studied. Tumors obtained from NOD/SCID mice xenografted with HuH7 and PLC/PRF/5 cells were digested to single cells and serially transplanted without isolation of cell-surface markers. BrdU was injected intraperitoneally. After 6 weeks for HuH7, and after 10 weeks for PLC/PRF/5, tumors were enucleated and sections were stained with anti-BrdU and anti-CD13 antibody. In the tumors derived from HuH7 cells, very small numbers of BrdU-retaining cells that also expressed CD13 were observed typically in the edge of tumor foci. BrdU-retaining cells were not observed in the tumor center. Tumors derived from PLC/PRF/5 cells grew more slowly than did those derived from HuH7 cells. BrdU-retaining cells could be identified and did express CD13. Interestingly, clusters of CD13+BrdU– cells were observed close to CD13+BrdU+ cells, suggesting that they might be derived from the CD13+BrdU+ cells (Figure D).
HCC-CD13+ cells form spheres and produce the CD90+ phenotype.
Sphere formation is a common characteristic of stem cells. To evaluate CD13 as a candidate CSC marker, the expression of CD13 in spheres derived from HuH7, PLC/PRF/5, and clinical HCC was studied. The expression of CD13 was increased in both HuH7 (2.0% in control vs. 67.0% in spheres; 33.5-fold increase) and PLC/PRF/5 (15.2% in control vs. 83.8% in spheres; 5.51-fold increase) (Figure A). There was no significant change in CD133 expression in HuH7. In PLC/PRF/5, expression of CD90 was decreased in the spheres (35.7% in control vs. 2.5% in spheres; 14.28-fold decrease). The expression of CD13 compared with that of CD133 and CD90 appeared to be associated with a more immature stem-like and dormant population. Spheres established from clinical HCC samples localized in the CD13+CD90–CD133– fraction in a manner similar to that observed in PLC/PRF/5 (Figure B).
CD13+ cells exist as a core in HCC spheres and produce CD90+ cells.
The time-course changes in the expression of CD13 and CD90 in PLC/PRF/5 were studied. Isolation and culture of the CD13+CD90– fraction from the PLC/PRF/5 spheres in serum-containing media resulted in the production of CD13+CD90+ fraction after 96 hours (Figure C). The isolated CD13–CD90– fraction elicited cell death within a few days and could not be maintained. Interestingly, the isolated CD13–CD90+ fraction rapidly produced the CD13+CD90– fraction within 24 hours (Figure C). These findings suggest that potentially dormant CD13+ cells produce proliferating CD90+ cells and that some parts of the proliferating CD90+ cells also produce CD13+ cells. It is important to determine how this CD13+ population (slow-growing potentially dormant) could be maintained in a cancer cell line in vitro. Dormant or slow-growing cell populations may disappear during continuous subculturing. These findings may replicate the rapid change from dormant to active status in cancer stem-like cells, as revealed by their cell-surface markers, or the dormant cells might mimic a certain multipotent condition in cellular differentiation.
CD13+ cells resist chemotherapy, and CD13 inhibition drives cells to apoptosis.
The change of cell-surface marker expression before and after doxorubicin (DXR) hydrochloride treatment or 5-fluorouracil (5-FU) was studied in HuH7 and PLC/PRF/5. In HuH7, CD13 expression was increased over 20-fold by DXR or 5-FU treatment compared with control (CD13+CD133+ population in control, 2.0%; by DXR treatment, 40.3%; by 5-FU treatment, 44.3%), although expression of CD133 remained unchanged (87.1% in control vs. 88.0% after DXR treatment, 88.7% after 5-FU treatment). In PLC/PRF/5, after treatment with DXR, the CD13+CD90– fraction was also increased and the CD13–CD90+ fraction was shifted to the CD13 positive (the CD13+CD90– fraction of control was 15.4% and of DXR treatment was 58.2%). After treatment with 5-FU, the remaining cells were more clearly localized in the CD13+CD90– fraction (77.8%) (Figure A). The chemo-resistance ability of the CD13+ cells was also confirmed by cell proliferation assay in HuH7. The CD13+CD133+ fraction was highly resistant to DXR compared with the CD13–CD133+ and CD13–CD133– fractions, indicating consistent changes in the markers following DXR treatment (Supplemental Figure 2A). Although the CD13–CD133– fraction exhibited slow cell growth in the proliferation and cell fate study (Figure , B and C), this fraction showed high chemosensitivity.
CD13+ cells resist chemotherapy, and inhibition of CD13 elicits cellular apoptosis.
Next, the effect of CD13 inhibition on cell proliferation in HuH7 was assessed. Cell proliferation was suppressed in a concentration-dependent manner after 72 hours exposure to the CD13-neutralizing antibody. At 10 and 20 μg/ml concentrations of the CD13-neutralizing antibody, cell proliferation was suppressed by approximately 80% at 24 hours and 95% at 72 hours (Figure B). The apoptosis assay showed that both the CD13-neutralizing antibody and CD13 inhibitor ubenimex induced apoptosis in both HuH7 and PLC/PRF/5 after 24 hours (Figure C). The CD13 antibody (clone WM15) has been shown to be specific to humans and to function as a neutralizing antibody (15
). Reportedly, ubenimex (bestatin) specifically blocks CD13, which antagonizes the zinc-binding site of the aminopeptidase N domain (16
). Ubenimex is used as a therapeutic agent for adult acute nonlymphatic leukemia (20
We then hypothesized that not only the ABC transporter (21
) but also CD13 is involved in cell protection against exposure to anticancer agents. DXR is a well-known ABC-transporter–dependent anticancer drug. We have established a DXR-resistant HuH7 clone in which 90% of cells survive in 0.5 μg/ml of DXR, whereas about 99% of parent HuH7 cells die at that concentration (Supplemental Figure 2, B and C). Inhibition of CD13 indicated approximately 50% suppression of cell proliferation in this clone (Figure D), and this finding suggests that CD13 inhibition can potentially suppress cells that may have multidrug-resistance capacities and remain viable after conventional anticancer drug treatments.
CD13 is expressed preferentially in therapy-resistant HCC cells.
To identify the expression of CD13 in clinical HCC, HCC samples were digested and hematopoietic Lin–CD45– fractions were further analyzed by multicolor flow cytometry. In all 12 clinical HCC samples, including 3 cases of non–hepatitis-derived HCC (1 case recurred after transcatheter arterial embolization [TAE]) and 9 cases of hepatitis-derived HCC (4 cases recurred after TAE), no CD133 expression was observed. In all cases, CD13 and CD90 expression was observed in the following 4 subpopulations: CD13+CD90+, CD13+CD90–, CD13–CD90+, and CD13–CD90–. In cases that recurred after TAE, the CD13+CD90– fraction was more abundant than that in non-TAE cases (48% ± 12% in TAE cases vs. 8% ± 4% in non-TAE cases; 6-fold increase), whereas the CD13–CD90+ fraction was more abundant in non-TAE cases than in TAE cases (40% ± 18% in non-TAE cases vs. 12% ± 5% in TAE cases; 3.3-fold increase) (Figure A). In all 12 clinical HCC samples, the expression patterns were very similar to that of PLC/RLF/5, indicating its usefulness as an HCC model. Of course, the percentages of cells just indicate the percentage that survived after mechanical and enzymatic digestion. The majorities of HCC cells retain the cellular functions of liver cells, accumulate fat and glycogen, and produce bilirubin. Also, they are relatively bigger than other kinds of cancer cells and may be more easily damaged by mechanical and enzymatic digestion.
CD13 expression in clinical HCC samples with or without TAE.
The expression of CD13 was confirmed in fresh frozen surgical specimens. The CD13+ HCC cells typically existed along the fibrous capsule forming cellular clusters after TAE. In non-TAE cases, the CD13+ HCC cells usually formed small cellular clusters inside the cancer foci (Figure B). CD13 was expressed on the cell surface in HCC cases. In normal liver samples, CD13 was expressed in the sinusoid with a linear staining pattern and in bile ducts with an intraductal pattern; this was different in the HCC samples. The immunohistochemical findings for the post-TAE cases support clinical experience because HCC recurrence after TAE usually occurs at the fibrous capsule and chemoresistant viable HCC cells exist mainly around the fibrous capsule.
Interestingly, some small canalicular structures near the bile ducts expressed CD13 on the cell surface, and these are suggested to be liver stem/progenitor cells, since it has been reported that normal liver stem/progenitor cells express CD13 (23
). In our studies, spheres established from the normal liver were predominantly CD13+
, with a multidifferentiation potential in both hepatocyte and cholangiocyte lineages (Supplemental Figure 3).
To assess whether the area of the fibrous capsule contributed to the maintenance of semiquiescent CD13+
cells, we stained fresh frozen tissues obtained from HCC parents with a hypoxia marker, carbonic anhydrase 9 (CA9) (24
). In hematopoietic stem cells, hypoxia is well known as a hypoxic niche that plays important roles in maintaining stem cells in a dormant phase (25
). In the TAE samples, expression of CA9 was localized along the fibrous capsule and coexpressed CD13. In the non-TAE samples, CA9+
cells formed cellular clusters in cancer foci and coexpressed CD13. In normal livers, CA9 expression was limited to the cell surface of bile ducts (Supplemental Figure 4).
CD13 inhibition elicits tumor regression.
For preliminary studies, HuH7 cells were transplanted into NOD/SCID mice and treated with 5-FU to determine whether the CD13+ fraction was enriched by treatment with a DNA synthesis inhibitor or not. We used 5-FU, the most common anticancer drug in HCC treatment, to simulate the clinical setting. After 3 days of intraperitoneal administration of 5-FU (30 mg/kg), most Ki67+ active cells were disrupted and remained only at small foci, and tumors were replaced by a majority of CD13+Ki67– cells. In the controls, CD13 expression was limited to a small fraction with cellular clustering, and most cells expressing CD13 were Ki67–. Conversely, in ubenimex-treated mice (20 mg/kg, 3 days), most of the CD13+ cells were disrupted and replaced with Ki67+ active cells (Figure A).
CD13 inhibition elicits cancer regression in vivo.
PLC/PRF/5 was then used for further analyses. The expression of markers in this cell line is similar to that in clinical HCC, and thus, the PLC/PRF/5 cell line is potentially useful as an HCC model. In control mice, CD13 expression was limited to a small fraction and most of the cells expressed CD90. After treatment with 5-FU (30 mg/kg, 5 days of injection and 2 days of withdrawal, 2 courses), most of the CD90+ cells were disrupted and tumors were replaced by a majority of CD13+ cells. After ubenimex treatment (20 mg/kg every day for 14 days), not only were many CD90+ cells present but CD13+ cells were also identified. Interestingly, in cases in which both ubenimex and 5-FU were administered, the majority of tumor cells were disrupted. We identified atypical, nonspecific CD13 expression in these cases (Figure B). Taken together with the findings that CD90+ cells produce CD13+ cells within 24 hours and that almost all of the CD13+ cells were disrupted by ubenimex plus 5-FU treatment, the CD13+ cells that appeared in the ubenimex treatment groups may have been newly produced from residual CD90+ cells. Costaining of Ki67 and CD13 revealed that CD13+ cells were negative for the expression of Ki67 (Supplemental Figure 5).
The highly deformed nuclei observed in the ubenimex–plus–5-FU treatment specimens suggested that DNA fragments were present. The DNA fragmentation status was thus assessed by in situ hybridization with terminal deoxynucleotidyl transferase (TdT). There were a few DNA fragments in both the control and 5-FU–treated specimens, whereas there were many more in the ubenimex-treated specimens. Especially in the specimens treated with ubenimex plus 5-FU, there were numerous DNA fragments in residual tumor cells (Figure B).
After 14 days of treatment, the tumor volume was significantly decreased in the ubenimex–plus–5-FU groups compared with the control and 5-FU or ubenimex groups (Figure , C and D).
Next, we studied the effects of CD13 inhibition as it pertains to the self-renewing ability of cells and repopulation of tumors. The CD13+-enriched fraction obtained from 5-FU–treated mice was serially transplanted into secondary NOD/SCID mice. Starting the day after transplantation, the mice were treated with ubenimex (20 mg/kg) for 7 days. After 3 weeks, no tumor formation was observed in the ubenimex-treated mice (n = 0/6), whereas 60% of the untreated mice grew tumors (n = 6/10) (Figure E).
The CD13+ HCC cells contain lower levels of ROS.
We focused on the ROS scavenger pathway to determine why DNA fragmentation and apoptosis were induced by CD13 inhibition. It has been reported that self-renewing dormant stem cells normally possess low levels of intracellular ROS and that deregulation of ROS levels impairs stem cell functions (27
). Intracellular ROS levels were measured by prooxidants using the 2′,7′-dichlorofluorescein diacetate (DCF-DA) stain. Both in HuH7 and PLC/PRF/5, the CD13+
fraction contained lower concentrations of ROS than the CD133strong
fractions. After stimulation of oxidative stress by H2
, a lower concentration of ROS was clearly observed in the CD13+
fraction compared with the CD13–
fraction. Following treatment with the CD13-neutralizing antibody or ubenimex, the ROS concentration was significantly increased in CD13+
cells and reached the level of ROS observed in the CD13–
fraction (Figure A). In clinical HCC samples, the results were similar to those in PLC/PRF/5, as the CD13+
fraction exhibited lower ROS levels than those in the CD13–
fractions (Figure B). The CD13+
fraction also contained another ROS indicator, MitoSOX (a highly selective marker for mitochondrial superoxide), which was markedly lower in the PLC and clinical HCC samples and less in HuH7 (Supplemental Figure 6).
CD13+ cells contain lower levels of ROS than CD13– cells.
To study the correlation between CD13 and the ROS scavenger pathway, the expression of Gclm was assessed by RT-PCR. Gclm encodes the glutamate-cysteine ligase that catalyses the rate-limiting synthesis step of glutathione (GSH), which works as a critical cellular reducing agent and anti-oxidant. Gclm was overexpressed in the CD13+CD90– fraction (P < 0.001) compared with the CD13+CD90+, CD13–CD90+, and CD13–CD90– fractions in PLC/PRF/5 and primary HCC cells (Figure C).
It is well known that cell destruction after exposure to cytotoxic chemotherapy and ionizing radiation is partially due to free radicals (28
). Given that the present study indicates a low ROS concentration in the CD13+
population, we were interested to see whether chemotherapy agents actually increase ROS level of CD13+
population. To study this, ROS levels of CD13+
populations in HuH7, and CD13+
populations in PLC/PRF/5 were measured 3 hours and 48 hours after of DXR or 5-FU treatment. After 3 hours treatment with DXR, ROS levels were increased in both CD13+
populations in HuH7 and PLC/PRF/5. Interestingly, after 48-hour treatment with DXR, ROS levels of CD13+
populations were decreased and reached those of control levels. Especially in PLC/PRF/5, CD13+
populations showed 2 peaks of ROS levels, one of which contained further lowered ROS levels than control. With 5-FU treatment, though the power of upregulation of ROS levels was weaker than those of DXR, ROS levels of CD13+
fractions were actually increased to those of CD13–
fractions. As with the data regarding DXR treatment, after 48 hours of 5-FU treatment, CD13+
populations showed lower levels of ROS compared with those of the CD13–
population (Figure D). These data together with the observation that CD13+
cells remained after treatment with chemotherapy agents (Figure A), suggest that ROS levels of all of the cells are temporally upregulated when cells are treated with chemotherapy agents and that this leads to disruption of the CD13–
population, whereas in the CD13+
cells, ROS levels are downregulated by the ROS scavenger pathway and the cells survive. In addition, proliferative CD13–
cells are easily affected by the DNA synthesis inhibition effect of chemotherapy agents.
To assess radiation-induced DNA damage with ROS, purified CD13+
, and CD13–
PLC/PRF/5 cells were irradiated and subjected to an alkaline comet assay. Although untreated cells did not show significantly different levels of DNA damage, there were fewer DNA strand breaks in CD13+
cells than in the other 3 fractions (P
< 0.01) after ionizing irradiation. The DNA damage in these 3 fractions (but not in the CD13+
fraction) was significantly inhibited (P
< 0.001) by treatment with an antioxidant reagent, tempol (Figure A). In HuH7 cells, the CD13+
fraction also exhibited lower levels of DNA damage compared with the CD13–
fraction. There was no significant difference between the irradiated and tempol-treated groups for the CD13–
fraction (Figure A). These findings reveal that the enhanced ROS defenses in the CD13+
fraction contribute to the reduction in DNA damage after genotoxic cancer therapy. To confirm radiation-induced DNA double-strand break status in CD13+
populations, time-course change of gamma-H2AX, a marker of double-strand breaks (30
), was studied. In PLC/PRF/5, after 4 Gy of irradiation, gamma-H2AX expression in CD13–
population increased after 2 hours of irradiation (45.4% ± 5.3%) and then decreased within 6 hours (15.6% ± 4.5%), whereas gamma-H2AX expression in CD13+
population did not. In HuH7, gamma-H2AX expression increased after 2 hours in both CD13+
populations and decreased rapidly in the CD13+
population (4.4% ± 2.8%) compared with the CD13–
population (38.4% ± 4.6%) (Figure B). After 24 hours of irradiation, the residual cells were localized in the CD13+
fraction in HuH7 and in the CD13+
fraction in PLC/PRF/5 (Figure C). Although there were some different manners in time-course change of gamma-H2AX in PLC/PRF/5 and HuH7, surviving cells after 24 hours of irradiation were localized in the CD13+
population, suggesting the radio-resistant characteristics of the CD13+
population, due to rapid recovery of DNA damage. After 48 hours of irradiation, the residual cells began to proliferate and produced CD13–
cells in HuH7 and CD13+
cells in PLC/PRF/5 (Figure C). These studies support the time-course studies (Figure C) and indicate that CD13+
cells exist as a core fraction in the cellular hierarchy.
High levels of ROS scavenger expression parallel DNA damage in CD13+ HCC cells.