Abstract
Background
Evaluating the non-clinical biodistribution (BD) of cell therapy products (CTPs) is essential to predicting their efficacy and safety in clinical trials. Although BD is typically quantified by qPCR following DNA extraction, this process is labor-intensive and time-consuming, and subject to variability in extraction yield, which complicates normalization to organ weight or blood volume. In contrast, direct qPCR enables the quantification of lysed samples without DNA extraction. Here, we systematically developed an effective direct qPCR method applicable to BD studies of CTPs.
Methods
We used HLCN061, a genetically modified natural killer (NK) cell derived from human induced pluripotent stem (iPS) cells. Mouse organs and blood containing HLCN061 cells were lysed, diluted, and directly subjected to real-time PCR using a human-specific Alu primer set. PCR reactions employed an inhibitor-tolerant master mix. Assay performance was evaluated by linearity (R2), amplification efficiency, relative error (RE), coefficient of variation (CV), and lower limit of quantification (LLOQ). The validated method was then applied to evaluate the BD of HLCN061 cells over 24 h following intravenous administration to mice.
Results
Across all tissues and blood samples, R2 exceeded 0.99, PCR amplification efficiency ranged from 90% to 110%, and RE was within ±25%. CV values were within 25% except for one case. LLOQ was 0.25 cells/mg weight for the liver, lungs, kidneys, and heart; 1 cell/mg weight for the spleen and ovaries; and 0.3 cells/μL for blood. Weight-normalized concentrations were highest in the lung, with the most rapid clearance from blood.
Conclusion
This direct qPCR method is effective and suitable for non-clinical BD assessment of CTPs.
Keywords: Biodistribution, Cell therapy products, Quantification, Direct qPCR, Validation
1. Introduction
Understanding the pharmacokinetics of cellular therapeutic products (CTPs) is critical to interpreting and predicting their efficacy and toxicity [1], [2]. To evaluate the biodistribution (BD) of CTPs, a method is used in which CTPs are administered to laboratory animals, followed by quantification of CTP-derived genes in organs and blood. The main quantitative method used to date is qPCR, primarily real-time PCR [3], [4], [5]. In recent years, droplet digital PCR has also been reported, which enables more precise quantification [6], [7], [8], [9].
qPCR-based evaluation typically involves DNA extraction from organs or blood, primarily using columns and magnetic beads. However, this process is complicated and time-consuming [1], [2], [10], [11], [12], [13]. Furthermore, because DNA extraction yields vary, the data obtained are based on genomic DNA quantity, limiting use in evaluations based on organ weight or blood volume [14]. This in turn hampers use in the evaluation of cellular kinetics, pharmacokinetics-pharmacodynamics (PK-PD), and quantitative modeling and simulation, because these analyses are based on volume or weight [8], [14]. In response, recent reports have noted that the addition of an external control gene to the sample and measurement of that specific gene allows for the correction of DNA extraction yield, and thereby enables evaluations based on organ weight or blood volume [7], [9], [14].
In contrast, direct qPCR, which quantifies samples obtained by lysing organs or blood directly via qPCR, does not require DNA extraction, enabling rapid and simple analysis [15], [16], [17], [18]. It also allows for the direct assessment of cell concentration per unit of organ weight or blood volume. To date, however, few studies have described the application of CTPs to non-clinical BD studies [19]. Furthermore, we are unaware of any published reports of the detailed quantitative sensitivity or validation of direct qPCR in non-clinical BD studies of CTPs. Moreover, since direct qPCR does not involve DNA extraction, the PCR reaction may be inhibited by surfactants in the lysis buffer used during lysis or by inhibitors present in the sample, potentially reducing sensitivity [15], [18].
In this study, we systematically developed for the first time an effective direct qPCR method for CTP biodistribution studies. The method used DirectAce qPCR mix (Nippon Gene, Tokyo, Japan) for the PCR reaction. The reagent suppresses the effects of PCR inhibitors, including surfactants used for cell lysis [20]. As human model cells, we used HLCN061, a gene engineered human induced pluripotent stem (iPS) cell-derived natural killer cell armed with NKG2D-DAP10 complex, IL-15, CD16 (F176V), CCL19, and CCR2B for the treatment of refractory solid tumors [21], [22]. qPCR targeting used the Alu sequence. Alu elements are primate-specific repetitive elements, with over 1 million copies present throughout the human genome [23], and the Alu sequence enables highly sensitive cell quantification. Under the experimental conditions, where HLCN061 cells were the only human cells administered or spiked into mouse matrices, it is reasonable to interpret the Alu signal as being derived from HLCN061 cells.
The direct qPCR method was evaluated in detail for calibration curve linearity, PCR amplification efficiency, quantitative sensitivity, accuracy, precision and reproducibility. Furthermore, the utility of the established direct qPCR method was demonstrated by evaluating the BD study of HLCN061 cells following intravenous administration to mice.
2. Materials and methods
2.1. Materials
HLCN061, human iPS-derived genetically modified NK cells, were established as described previously [21]. Briefly, human iPSCs derived from umbilical cord blood CD34+ cells were genetically modified to introduce six transgenes (NKG2D-DAP 10 complex, IL-15, CD16 (F176V), CCL19, and CCR2B) and subsequently differentiated into NK cells.
Control mouse blood and tissues (liver, kidney, heart, lung, spleen, and ovary) were obtained from female NOD.Cg-Prkdcscid Il2rgtm1Sug/ShiJic (NOG) mice (In-Vivo Science International Inc., Kawasaki, Japan). These mice were housed in animal facilities under specific pathogen-free (SPF) conditions with ad libitum access to water and food. All animal protocols were performed in accordance with the relevant institutional and national guidelines and regulations and were approved by the Institutional Animal Care and Use Committee of the HEALIOS K.K.
Primers and probes for the human Alu sequence were prepared by Integrated DNA Technologies, Inc. (Coralville, IA, USA). The sequences are as follows: Primers: 5′-CAT GGT GAA ACC CCG TCT CTA-3′ (forward) and 5′-GCC TCA GCC TCC CGA GTA G-3′ (reverse); Probe: 5′-ATT AGC CGG GCG TGG TGG CG-3′ [24]. The probe contains the reporter fluorophore FAM at the 5′ end, an internal ZEN quencher, and an Iowa Black FQ quencher at the 3′ end. The expected amplicon size was approximately 90 bp.
2.2. Development of the direct qPCR method
Fig. 1 shows the procedure used for the direct qPCR method. The concentrations of HLCN061 cells in liver, kidney, heart, lung, spleen, ovary and blood of female NOG mice were estimated. HLCN061 cells lysate was prepared by adding MagMAX™ Cell and Tissue DNA Extraction Buffer (Thermo Fisher Scientific, Waltham, MA, USA). Standard samples for the HLCN061 cells calibration curve were prepared by diluting the cell lysate with MagMAX™ Cell and Tissue DNA Extraction Buffer.
Fig. 1.
Procedure of the direct qPCR method.
Control mouse liver, kidney, heart, lung, spleen, and ovary tissues were homogenized in PBS (Thermo Fisher Scientific) at a concentration of 20 mg tissue per 100 μL. For each tissue homogenate, HLCN061 cells calibration curve standard sample and MagMAX™ Cell and Tissue DNA Extraction Buffer were added, together with MagMAX™ DNA Multi-Sample Ultra 2.0 Enhancer Solution and MagMAX™ DNA Multi-Sample Ultra 2.0 Proteinase K (Thermo Fisher Scientific). The volume ratio of the solution containing tissue homogenate and HLCN061 cells calibration standard sample added to MagMAX™ Cell and Tissue DNA Extraction Buffer, MagMAX™ DNA Multi-Sample Ultra 2.0 Enhancer Solution, and MagMAX™ DNA Multi-Sample Ultra 2.0 Proteinase K was set to 500:20:40. For blood samples, 50 μL of whole blood was mixed with 450 μL of HLCN061 cells calibration curve standard sample and MagMAX™ Cell and Tissue DNA Extraction Buffer, together with 20 μL of Enhancer Solution and 40 μL of Proteinase K under the same conditions.
The samples were incubated at 65°C for 2 h to prepare the crude samples. The term ‘crude sample volume’ refers to the total volume of the crude lysate prepared prior to qPCR dilution, after mixing the biological matrix (tissue homogenate or blood) with the HLCN061 cells standard solution, MagMAX™ Cell and Tissue DNA Extraction Buffer, MagMAX™ DNA Multi-Sample Ultra 2.0 Enhancer Solution, and MagMAX™ DNA Multi-Sample Ultra 2.0 Proteinase K. The prepared crude samples were diluted 3–100-fold with distilled water (Thermo Fisher Scientific).
Real-time PCR was performed using a total reaction volume of 30 μL, consisting of 12 μL of diluted crude sample, 15 μL of DirectAce qPCR mix or PrimeTime Gene Expression Master Mix (Integrated DNA Technologies, Inc.), 1.08 μL each of forward and reverse primer solutions (25 μM), 0.6 μL of probe solution (10 μM), and 0.24 μL of distilled water (Thermo Fisher Scientific).
Real-time PCR assays were performed using the QuantStudio 5 Flex Real Time PCR system equipped with QuantStudio Design & Analysis Software (Thermo Fisher Scientific). The PCR conditions were as follows: after incubation at 95°C for 10 min (DirectAce qPCR mix) or 3 min (PrimeTime Gene Expression Master Mix), 40 cycles were performed at 95°C for 15 s followed by 60°C or 61°C for 60 s. The Ct cutoff criterion for detection was set at 40 cycles. Samples with no detectable amplification (Ct > 40 or undetermined) were classified as non-detectable (ND).
Calibration standards were subjected to the same crude sample preparation, dilution, and qPCR procedures as the study samples. Therefore, the tissue weight or blood volume, crude sample preparation process, dilution factor, and qPCR input volume were inherently incorporated into the calibration curves.
The concentration of HLCN061 cells was calculated using equation X = 10 (Y−b)/a, derived from the calibration curve. Here, X represents the calculated cell concentration (cells/mg tissue or cells/μL blood), Y represents the cycle threshold (Ct) value of each sample, and a and b represent the slope and y-intercept of the calibration curve, respectively. For example, when the Ct value of a liver sample was substituted into the above equation, the calculated value directly represented the concentration of HLCN061 cells in liver tissue (cells/mg liver), because the calibration standards and study samples underwent identical preparation and analytical procedures. The obtained values (cells/mg tissue or cells/μL blood) represent the HLCN061 cell equivalents, based on a calibration curve derived from HLCN061.
PCR efficiency (E) was calculated using equation E = 10 (−1/slope) - 1.
Fig. 2 shows the experimental condition setting of the direct qPCR method. The experimental conditions described below for the direct qPCR method were established to satisfy the following criteria: for the calibration curve, linearity (R2 ≥ 0.99) and PCR amplification efficiency (90–110%).
Fig. 2.
Experimental condition setting of the direct qPCR method.
Next, using Quality Control (QC) samples, the method was evaluated against the following criteria: relative error (RE) ≤ |±25|% and coefficient of variation (CV) ≤ 25%. Each of the High Quality Control (HQC), Medium Quality Control (MQC), Low Quality Control (LQC), and Lower Limit Quality Control (LLQC) (=Lower Limit of Quantification (LLOQ)) samples was tested in quadruplicate (n = 4). The concentration range of the calibration standards was determined based on preliminary optimization tests and the expected concentration range of HLCN061 cells in in vivo study. The HQC concentration was set at 80% of the HLOQ. The MQC concentration was arbitrarily chosen to fall between the HQC and LQC levels, while ensuring it represents a different order of magnitude. The LQC concentration was defined as 2–3 times the LLOQ. The LLQC concentration was set equivalent to the LLOQ. The LLOQ concentrations were set as concentrations that met the RE and CV criteria on a calibration curve where linearity and PCR amplification efficiency satisfied the established criteria. This analysis was designated as Run 1. Table 1 shows the nominal concentrations of the standard and the QC samples in each organ and in blood.
Table 1.
Nominal concentrations of standard and QC samples.
| (a) liver, kidney, heart, lung | |
|---|---|
| Nominal concentration (cells/mg tissue) | Standard or QC |
| 500 | S1 |
| 400 | HQC |
| 150 | S2 |
| 50 | S3 |
| 40 | MQC |
| 15 | S4 |
| 5 | S5 |
| 0.75 | LQC |
| 0.5 | S6 |
| 0.25 | LLOQ, LLQC |
| (b) spleen, ovary | |
| Nominal concentration (cells/mg tissue) | Standard or QC |
| 1000 | S1 |
| 800 | HQC |
| 300 | S2 |
| 100 | S3 |
| 80 | MQC |
| 30 | S4 |
| 10 | S5 |
| 3 | S6, LQC |
| 1 | LLOQ, LLQC |
| (c) blood | |
| Nominal concentration (cells/μL blood) | Standard or QC |
| 200 | S1 |
| 160 | HQC |
| 60 | S2 |
| 20 | S3 |
| 16 | MQC |
| 6 | S4 |
| 2 | S5 |
| 0.6 | S6, LQC |
| 0.3 | LLOQ, LLQC |
S1-S6, LLOQ: standard.
HQC, MQC, LQC, LLQC: QC.
To establish concentration measurement methods for each organ and blood that met the above criteria, the following experimental conditions were optimized: organ weight or blood volume used for analysis, crude sample volume, LLOQ, dilution factor of crude samples, and annealing temperature.
To confirm that direct qPCR specifically quantified HLCN061 cells, measured values were compared between blank and lower limit quality control (LLQC) samples (n = 4) for each organ and blood matrix.
To confirm the selectivity of direct qPCR for HLCN061 cells in biological matrices, measured values were compared between blank and LLOQ-spiked samples prepared from blood collected from six NOG mice. Each blood sample was divided into two aliquots: a blank sample (unspiked blood) and an LLOQ-spiked sample (blood spiked with HLCN061 cells at the LLOQ).
2.3. Method validation
In addition to Run 1, described in Section 1.2. Above, Runs 2 and 3 were conducted using the same method to determine concentrations in various organs and blood to confirm reproducibility (Fig. 2). QC samples were analyzed in quadruplicate (n = 4).
2.4. In vivo study
Female NOG mice were housed as described in Section 1.1. All animal protocols were performed in accordance with the relevant institutional and national guidelines and regulations and were approved by the Institutional Animal Care and Use Committee of the HEALIOS K.K.
HLCN061 cells were administered intravenously to female NOG mice (n = 3) at a dose of 5 × 106 cells/mouse. Blood and organs (liver, kidney, heart, lung, spleen, and ovary) were collected at 5 min, 1 h, and 24 h after administration. The samples were immediately placed on ice and stored at −80°C until analysis.
3. Results
3.1. Development of the direct qPCR method
We compared calibration curves, R2 and PCR amplification efficiencies for liver crude samples diluted 3-, 10-, 30-, and 100-fold using both DirectAce qPCR mix and a conventional master mix (Fig. 3). The results demonstrated that amplification with DirectAce qPCR mix was observed starting at a 3-fold dilution. In contrast, no amplification was observed with the conventional master mix at 3- or 10-fold dilutions, with amplification detected only starting at a 30-fold dilution. Furthermore, the R2 value and PCR amplification efficiency of the calibration curve at a 10-fold dilution with DirectAce qPCR mix were 0.998 and 101.8%, respectively, meeting the criteria of R2 ≥ 0.99 and the PCR amplification efficiency within 90–110%. In contrast, the conventional master mix achieved an R2 value of 0.999 and PCR amplification efficiency of 106.9% only at a 100-fold dilution.
Fig. 3.
Effects of dilution ratios on calibration curves, R2 and PCR amplification efficiencies in liver sample
Liver crude samples were diluted 3-, 10-, 30-, and 100-fold with distilled water and analyzed by qPCR using DirectAce qPCR mix and a conventional master mix. Calibration curves were generated for each dilution ratio. The R2 values and PCR amplification efficiency were calculated from the calibration curves and compared between the two qPCR master mixes.
Table 2 summarizes the experimental conditions that met the established criteria. Each experimental condition had to be optimized for each organ and blood concentration measurement. The R2 values, PCR amplification efficiency, and RE and CV values at each QC concentration for Run 1 are shown in Fig. 4. All values met the criteria. To further validate the method, Ct values for LLOQ and blank samples are provided in Table S1.
Table 2.
Conditions of the direct qPCR method in tissue and blood samples.
| Liver | Kidney | Heart | Lung | Spleen | Ovary | Blood | |
|---|---|---|---|---|---|---|---|
| mg of organ or μL blood/sample | 20 | 20 | 10 | 10 | 6 | 5 | 50 |
| Tissue homogenate or blood volume (μL) | 100 | 100 | 50 | 50 | 30 | 25 | 50 |
| HLCN061 cells standard solution volume (μL) | 50 | 50 | 25 | 25 | 30 | 25 | 50 |
| MagMAX™ Cell and Tissue DNA Extraction Buffer volume (μL) | 350 | 350 | 175 | 175 | 240 | 200 | 400 |
| MagMAX™ DNA Multi-Sample Ultra 2.0 Enhancer Solution volume (μL) | 20 | 20 | 10 | 10 | 12 | 10 | 20 |
| MagMAX™ DNA Multi-Sample Ultra 2.0 Proteinase K volume (μL) | 40 | 40 | 20 | 20 | 24 | 20 | 40 |
| Crude sample volume (μL) | 560 | 560 | 280 | 280 | 336 | 280 | 560 |
| LLOQ (cells/mg tissue or μL blood) | 0.25 | 0.25 | 0.25 | 0.25 | 1 | 1 | 0.3 |
| HLCN061 cells per PCR reaction at LLOQ (cells/reaction) | 0.011 | 0.011 | 0.011 | 0.0054 | 0.0071 | 0.0071 | 0.0054 |
| Tissue amount or blood volume per PCR reaction (mg tissue or μL blood/reaction) | 0.043 | 0.043 | 0.043 | 0.021 | 0.0071 | 0.0071 | 0.018 |
| Dilution ratio of crude sample | 10 | 10 | 10 | 20 | 30 | 30 | 60 |
| Annealing temperature (°C) | 60 | 60 | 60 | 61 | 60 | 60 | 60 |
Crude sample volume: Total volume of the crude lysate prepared prior to qPCR dilution, after mixing the biological matrix (tissue homogenate or blood) with the HLCN061 cells standard solution, MagMAX™ Cell and Tissue DNA Extraction Buffer, MagMAX™ DNA Multi-Sample Ultra 2.0 Enhancer Solution, and MagMAX™ DNA Multi-Sample Ultra 2.0 Proteinase K.
Fig. 4.
Validation of the direct qPCR method in tissue and blood samples, The R2 values, PCR amplification efficiencies calculated from calibration curves and the RE and CV values for QC samples obtained in Runs 1, 2, and 3 are presented. Bars represent the RE (%) and CV (%) for each QC sample. Each of QC sample was tested in quadruplicate (n = 4). (.): nominal concentrations of HQC, MQC, LQC and LLQC (cells/mg tissue or μL blood)
The overall distributions and numerical data of RE and CV values are shown in Fig. S1 and Table S2, respectively.
To evaluate specificity, we compared blank and LLQC samples in each organ and in blood (Table 3). The LLQC samples showed concentrations consistent with the nominal values, and the RE and CV values met the criteria. The blank relative to the LLOQ ranged from 6.8% for ovary to 24.8% for lung, and the blank samples showed clearly lower apparent concentrations, confirming the validity of the selected LLOQ levels.
Table 3.
Concentrations of human cells in blank and HLCN061 cells-spiked samples.
| Nominal concentration at LLOQ (cells/mg tissue or μL blood) | Blank (cells/mg tissue or μL blood) | LLOQ (cells/mg tissue or μL blood) | Mean ± SD (cells/mg tissue or μL blood | RE (%) | CV (%) | Ratio of blank to LLOQ samples (%) | |
|---|---|---|---|---|---|---|---|
| Liver | 0.25 | 0.043 | 0.24, 0.25, 0.25, 0.25 | 0.25 ±0.0028 |
−1.19 | 1.13 | 17.2-17.9 |
| Kidney | 0.034 | 0.28, 0.35, 0.26, 0.28 | 0.29 ±0.038 |
17.5 | 12.8 | 9.71-13.1 | |
| Heart | 0.036 | 0.28, 0.27, 0.29, 0.25 | 0.27 ±0.016 |
9.77 | 5.72 | 12.4-14.4 | |
| Lung | 0.062 | 0.30, 0.36, 0.25, 0.27 | 0.29 ±0.047 |
16.9 | 16.1 | 17.2-24.8 | |
| Spleen | 1 | 0.11 | 1.0, 1.1, 1.1, 0.94 | 1.1 ±0.079 |
5.22 | 7.54 | 10.0-11.7 |
| Ovary | 0.082 | 1.2, 1.2, 1.1, 1.1 | 1.1 ±0.074 |
14.3 | 6.47 | 6.83-7.45 | |
| Blood | 0.3 | 0.049 | 0.25, 0.31, 0.32, 0.34 | 0.31 ±0.041 |
2.09 | 13.3 | 14.4-19.6 |
To evaluate selectivity, we compared blank and LLOQ samples using blood collected from six NOG mice (Table 4). The blank relative to the LLOQ ranged from 19.1% to 33.7%. In all samples, the measured concentrations in the blank samples were clearly lower than those in the LLOQ samples, confirming the validity of the selected LLOQ levels. Furthermore, RE at the LLOQ was 16.0% and the CV was 17.2%, both of which met the criteria.
Table 4.
Concentrations of human cells in blank and HLCN061 cells-spiked individual mouse blood samples.
| Nominal concentration at LLOQ (cells/μL blood) | Blank (cells/μL blood) | LLOQ (cells/μL blood) | Ratio of blank to LLOQ samples (%) | |
|---|---|---|---|---|
| Mouse No. 1 | 0.3 | 0.066 | 0.31 | 21.3 |
| Mouse No. 2 | 0.091 | 0.27 | 33.7 | |
| Mouse No. 3 | 0.080 | 0.39 | 20.5 | |
| Mouse No. 4 | 0.080 | 0.34 | 23.5 | |
| Mouse No. 5 | 0.084 | 0.33 | 25.5 | |
| Mouse No. 6 | 0.084 | 0.44 | 19.1 | |
| Mean (cells/μL) | 0.081 | 0.35 | 23.1 | |
| RE (%) | - | 16.0 | ||
| CV (%) | 10.3 | 17.2 |
3.2. Method validation
In addition to Run 1, Fig. 4 shows the R2 values and PCR amplification efficiencies for the calibration curves in Runs 2 and 3, as well as the RE and CV values for each QC sample. All R2 values and PCR efficiencies met the acceptance criteria (R2 ≥ 0.99 and 90% ≤ PCR efficiency ≤ 110%). Except for a single value (heart, Run 3 LLQC, CV = 25.03%), all RE and CV values were within the acceptable range (−25% to 25% for RE and ≤ 25% for CV). The numerical data of RE and CV values are shown in Table S2. Of all RE values, 88.1% fell within ±20%, and 71.4% fell within ±15%. Furthermore, of all CV values, 96.4% fell within 20%, and 88.1% fell within 15% (Fig. S1).
3.3. In vivo study
Fig. 5 shows the concentrations of HLCN061 cells in organs and blood following intravenous administration to female NOG mice. The lungs exhibited the highest concentrations from 5 min to 24 h. Clearance in the spleen and kidneys tended to be slower than in other organs. Clearance from blood was faster than from any organ.
Fig. 5.
Time-course of HLCN061 cell concentrations in tissues and blood of female NOG mice following a single intravenous administration of 5 × 106 cells/mouse. Data are presented as mean ± S.D. (n = 3 individual animals) for liver, kidney, heart, lung, spleen, and blood and are expressed as cells/mg tissue for organ samples (a) and cells/μL for blood samples (b). Ovary samples in (a) were prepared by pooling tissues from three animals at each time point and were analyzed as a single sample; therefore, only a single value (cells/mg tissue) is shown. Cell concentrations were determined using the direct qPCR method.
4. Discussion
In this study of an effective direct qPCR method for BD studies of CTPs, we found that R2 exceeded 0.99, PCR amplification efficiency ranged from 90% to 110%, and RE was within ±25% across all tissues and blood samples. Further, CV values were within 25% except for one case. LLOQ was 0.25 cells/mg weight for the liver, lungs, kidneys, and heart; 1 cell/mg weight for the spleen and ovaries; and 0.3 cells/μL for blood. These findings indicate that this direct qPCR method is effective and feasible for non-clinical BD assessment of CTPs. Although LLOQ values varied among matrices when expressed as cells/mg tissue, the cell equivalents per PCR reaction at the LLOQ were comparable across matrices (0.0054-0.011 cells/reaction) (Table 2). This result suggests that the sensitivity of the qPCR assay was comparable across the different matrices. In contrast, the amount of tissue analyzed per PCR reaction ranged from 0.0071 to 0.043 mg tissue/PCR reaction (Table 2) and did not follow the same pattern as the cell equivalents per PCR reaction at the LLOQ. Therefore, the observed differences in LLOQ (cells/mg tissue) were mainly due to the amount of tissue represented in each PCR reaction, rather than differences in intrinsic assay performance.
Of note, the results showed that the selection of the master mix was critical for sensitivity. DirectAce qPCR mix met the calibration criteria at substantially lower dilution ratios than a conventional master mix. Although direct qPCR is a simple method that does not require DNA extraction [15], [16], [17], [18], the PCR reaction may be inhibited by surfactants in the lysis buffer or by inhibitors present in the sample [15], [18]. Of note, DirectAce qPCR mix suppresses this PCR inhibition [20]. With the conventional master mix, R2 was 0.999 at a 100-fold dilution, and the amplification efficiency was 106.9%, meeting the criteria (Fig. 3). On re-examination under the same conditions, however, R2 was 0.985 and amplification efficiency was 134.7%, which failed to meet the criteria, and reproducibility was not observed (data not shown). These findings indicated that use of the DirectAce qPCR mix as master mix improved quantitative performance.
The reagents used for sample lysis and digestion are critical for establishing sensitive direct qPCR. In this study, MagMAX™ Cell and Tissue DNA Extraction Buffer, MagMAX™ DNA Multi-Sample Ultra 2.0 Enhancer Solution, and MagMAX™ DNA Multi-Sample Ultra 2.0 Proteinase K were utilized. These reagents are typically employed for DNA extraction using magnetic bead-based methods.
To establish the direct qPCR method, we first defined acceptance criteria for the calibration curve R2, PCR amplification efficiency, and QC. Among these criteria, we set “linearity of the calibration curve: R2 ≥ 0.99” and “90% ≤ PCR amplification efficiency ≤ 110%.” Several studies have proposed similar criteria (R2 ≥ 0.98–0.99 and 90%–110% PCR amplification efficiency) [2], [12], [25], [26], [27].
On the other hand, for the criteria for RE and CV, we set −25% ≤ RE ≤ 25% and CV ≤ 25%. Although no guidelines for these values have yet been established [7], [8], previous studies have proposed various criteria, with reported CV thresholds ranging from less than 25% to less than 80% [8], [12], [25], [26]. Accordingly, the criteria used in this study are comparatively stringent.
To meet these criteria, it was necessary to optimize different experimental conditions for each organ and blood concentration measurement, including the amount of tissue or blood used, crude sample volume, LLOQ, dilution factor of the crude sample, and annealing temperature. In addition, the amount of tissue used for each organ was determined based on the total organ weight (Table 2). While the direct qPCR method requires matrix-specific optimization during the initial development phase, it is important to note that this represents a one-time effort. Routine analysis can be performed using the established conditions for each tissue matrix.
Regarding the specificity of the direct qPCR method, blank samples showed lower apparent concentrations than LLOQ samples across all organs and blood (Table 3). These observations support the validity of the selected LLOQ levels. These results indicate that the established direct qPCR method specifically detects HLCN061 cells. Additionally, the measured concentrations of LLQC samples are consistent with nominal LLOQ values, and the RE and CV values met the criteria. These results demonstrate the reliability of the method. The Ct values for blank and LLOQ samples also provide additional evidence of the reliability of the method (Table S1).
Regarding selectivity, the direct qPCR method reliably distinguished HLCN061 cells from background across six individual mouse blood matrices, and blank values were clearly separated from LLOQ values in every matrix (Table 4). The RE and CV at the LLOQ, 16.0% and 17.2%, were within the limits set for method validation. The method therefore quantifies HLCN061 cells accurately even in the presence of other mouse matrix components.
The specificity and selectivity of the direct qPCR method were evaluated by comparing blank and LLOQ samples, which supports the validity of the selected LLOQ levels. However, only a single blank measurement was available for each tissue matrix. Therefore, the distribution and variability of background signals could not be evaluated, and statistical separation between blank and LLOQ values was not assessed. Additional studies with larger numbers of blank samples may further characterize background signal variability and support more rigorous evaluation of assay specificity and selectivity.
In the validation conducted for Runs 1, 2, and 3, all R2 values exceeded 0.99, and amplification efficiencies fell within the acceptable range of 90–110% on the calibration curves. Furthermore, with the exception of the CV value for the heart sample in Run 3 LLQC, all RE and CV values met the criteria. The CV value for the heart sample in Run 3 LLQC was 25.03%, slightly exceeding the criterion of CV ≤ 25% (Fig. 4). Importantly, this minor exception did not clearly affect the overall precision or accuracy of the method. Furthermore, 88.1% of all RE values fell within a range of ±20%, and 96.4% of all CV values fell within a range of 20%, indicating favorable results (Fig. S1). These findings demonstrate that the direct qPCR method showed acceptable precision, accuracy, and reproducibility.
A key advantage of direct qPCR is that it expresses cell number per unit organ weight or blood volume — the form required to construct physiological and other pharmacokinetic models of CTP distribution. By contrast, the variable extraction yields of DNA-extraction qPCR produce data based on gDNA amount, which markedly hampers evaluation on a weight or volume basis [14].
Nakayama et al. recently reported a measurement method using droplet digital PCR with a primer set which targets the retrotransposon LINE-1, with human peripheral blood-derived CD8+ T cells as the target [7]. As an external control gene, canine genomic DNA was added to the samples prior to DNA extraction. The authors demonstrated that canine-specific qPCR performed during this process enabled correction for differences in DNA recovery rate and allowed evaluation in terms of cell number per organ weight or blood volume. On comparison, our present direct qPCR method reported LLOQs for the liver, kidney, and heart of 0.25 cells/mg, whereas Nakayama et al. reported a comparable value of 0.15 cells/mg. In contrast, the LLOQs obtained by direct qPCR for the spleen and blood were 1 cell/mg spleen and 0.3 cells/μL blood, respectively, or approximately three-fold (spleen) and five-fold (blood) higher than those reported by Nakayama et al.
Regarding precision and accuracy, although Nakayama et al. reported RE and CV values which exceeded 30% in some cases, most values were within 25%, indicating performance comparable to that of the direct qPCR method.
Although the LLOQs for spleen and blood were higher with direct qPCR, the differences were limited to approximately three-to five-fold.
The principal practical advantage of direct qPCR is that it eliminates DNA extraction and subsequent DNA quantification steps, and external control gene-based correction procedures, thereby simplifying the analytical workflow. These advantages remain relevant even when automated DNA extraction systems are available, because the extraction process and its associated reagents are still required. On the other hand, implementation of the method requires matrix-specific optimization of experimental conditions, including tissue amount, crude sample volume, dilution factor, and annealing temperature. Therefore, while the initial method development requires optimization, routine analysis can be performed using the established conditions for each tissue matrix. Taken together, the direct qPCR method employed in this study provides a practical alternative to DNA extraction-based approaches while maintaining comparable analytical performance.
Given its simplicity, precision, accuracy, and reproducibility, the direct qPCR method employed in this study can be broadly considered comparable to previously reported methods involving DNA extraction and the use of external control genes, although no direct head-to-head comparisons were performed in the present study.
The choice of target sequence and primer set warrants further optimization. The method described by Nakayama et al. uses LINE-1 as the target gene [7], [9], while Fujita et al. compared multiple primer sets for Alu elements [8]. Future studies should evaluate additional Alu primer sets, as well as primers which target non-Alu sequences.
With regard to PCR methods, this study employed real-time PCR. In contrast, Nakayama et al. and Fujita et al. investigated quantitative methods using digital PCR [7], [8], [9]. It has been reported that digital PCR offers superior accuracy and reproducibility compared to real-time PCR [28], [29]. The application of digital PCR to the direct qPCR approach also remains a future challenge. It is believed that such an approach will enable the establishment of more sensitive direct qPCR.
However, since real-time PCR generally offers a broader dynamic range than digital PCR [30], it is considered appropriate to use real-time PCR for high-concentration samples and digital PCR for low-concentration samples.
Using the direct qPCR method established in this study, we evaluated the BD of HLCN061 cells up to 24 h after intravenous administration. The lungs showed the highest levels from 5 min to 24 h. The concentration in blood declined more rapidly than in any organ.
Direct qPCR allows for the quantification of cell numbers per unit of organ weight or blood volume, enabling such appropriate comparisons between organs and blood. The present findings suggest that direct qPCR may provide useful information for the BD characteristics of CTPs. However, the applicability of this approach to other CTPs remains to be investigated.
On the other hand, apparent background concentrations were observed in blank samples (Table 3, Table 4). However, since untreated or vehicle-treated control animals were not included in the biodistribution study, the background signal level in vivo was not directly evaluated under the conditions of this study. Therefore, measurements obtained near the LLOQ or during the later stages of the assay may be affected by this background signal and should be interpreted with caution.
BD studies following NK cell administration to date have primarily reported cell concentration per total DNA as the concentration in organs and blood [31]; however, the present results suggest that direct qPCR may provide a valid assessment for pharmacokinetic studies.
Three limitations of this study warrant mention. First, the direct qPCR method required that experimental conditions be optimized separately for each organ and for blood concentration measurements (Table 2). We expect that further clarification of the differences in experimental conditions among organs and blood will allow the more systematic and efficient establishment of quantitative methods. Second, we used HLCN061 cells and NOG mice to evaluate the direct qPCR method. Further studies are needed to confirm whether the direct qPCR method is also useful in other cell types and mice. Third, we evaluated the applicability of this method to non-clinical BD studies for CTPs. Application of this method to other new modalities, such as viral therapies and nucleic acid-based therapeutics, is expected.
5. Conclusion
In this study, we systematically developed a method for quantifying cell concentrations in organs and blood using direct qPCR, designed for non-clinical BD studies of CTPs. We used DirectAce qPCR mix, which suppresses PCR inhibition. This master mix provided acceptable quantitative sensitivity, RE, CV, and reproducibility in the concentration measurements for each organ and in blood. These findings demonstrate the feasibility of the direct qPCR method in non-clinical BD studies of CTPs.
Author contributions
Y.N.: Conceptualization, Performed the experiments, Methodology, Data curation, Data analysis, Visualization, Writing – original draft, review & editing, Y.S.: Performed the experiments, Writing– review & editing, K.G., Y.T.: Writing – review & editing, H.K., K.T.: Writing – review & editing, Funding acquisition.
All authors have read and agreed to the published version of the manuscript.
Ethics declaration
Written informed consent to take part in the study and to publish the article has been obtained from all participants or their legal representatives. The privacy rights of participants have been observed.
This study was performed in compliance with relevant laws, regulatory frameworks and guidelines where the research took place. This study was approved by the committee of ethics on experimentation in HEALIOS K.K. (Approval No. C10091)
This study was conducted in accordance with the ARRIVE (Animal Research: Reporting of In Vivo Experiments) guidelines. This study was approved by the Institutional Animal Care and Use Committee of the HEALIOS K.K. (Approval No. 1234)
Declaration of generative AI and AI-assisted technologies in the writing process
During the preparation of this work, the authors used HEROZ ASK and ChatGPT to improve clarity and readability. After using these tools, the authors reviewed and edited the manuscript and take full responsibility for the published article.
Funding
This study was supported by Akatsuki Therapeutics Inc. And the Japan Agency for Medical Research and Development (AMED) under grant number JP24be0904017.
Declaration of competing interest
The authors declare the following financial interests/personal relationships which may be considered as potential competing interests: Y.N., Y.S., K.G., and Y.T. are employees of HEALIOS K.K.; H.K. is an executive officer of HEALIOS K.K.; K.T. is a senior advisor of HEALIOS K.K.
Footnotes
Peer review under responsibility of the Japanese Society for Regenerative Medicine.
Supplementary data to this article can be found online at https://doi.org/10.1016/j.reth.2026.101189.
Appendix A. Supplementary data
The following is the Supplementary data to this article:
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