Development of a GMP-Compliant Separation Method for Isolating Wharton's Jelly Derived Mesenchymal Stromal Cells from the Umbilical Cord

doi:10.21203/rs.3.rs-3326244/v1

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Research Article

Development of a GMP-Compliant Separation Method for Isolating Wharton's Jelly Derived Mesenchymal Stromal Cells from the Umbilical Cord

https://doi.org/10.21203/rs.3.rs-3326244/v1

This work is licensed under a CC BY 4.0 License

Journal Publication

published 03 May, 2024

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Background

Wharton's jelly derived mesenchymal stem cells (UC-MSCs) hold great therapeutic potential in regenerative medicine. However, GMP-compliant optimal methods for isolating UC-MSCs from UC tissue are still lacking. Additionally, there is a dearth of detailed research spanning from laboratory-scale to pilot-scale studies. Therefore, it is essential to establish standardized protocols that ensure the reproducibility and safety of UC-MSC manufacturing.

Methods

In this study, we aimed to explore and optimize parameters for the enzymatic digestion method used for isolating UC-MSCs. These parameters included enzyme concentrations, digestion times, seeding densities, and culture media. Additionally, we conducted a comparative analysis between the explant method and enzymatic digestion method. Subsequently, we evaluated the consecutive passaging stability of UC-MSCs, specifically up to passage 9, using the optimized enzymatic digestion method. Finally, we developed and scaled up manufacturing processes, starting from laboratory-scale flask-based production and progressing to pilot-scale cell factory-based production.

Results

The optimal parameters for the enzymatic digestion method were determined to be a concentration of 0.4 PZ U/mL Collagenase NB6 and a digestion time of 3 hours, resulting in a higher quantity of P0 UC-MSCs. Additionally, we observed a positive correlation between the initial cell seeding density and the number of P0 UC-MSCs. Evaluation of different concentrations of human platelet lysate (hPL) revealed that 2% and 5% concentrations resulted in similar levels of cell expansion, whereas a 10% concentration led to decreased cell expansion. Comparative analysis revealed that the enzymatic digestion method exhibited faster outgrowth of UC-MSCs compared to the explant method. However, after subsequent passages, there were no significant differences between the explant and enzymatic digestion methods in terms of cell proliferation, cell viability, and immunophenotype. Notably, consecutive passaging of UC-MSCs using the enzymatic digestion method demonstrated stability, with maintained cellular characteristics and functionality. Passages 2 to 5 exhibited higher viability and proliferation ability. Moreover, we successfully developed scalable manufacturing processes from the laboratory scale to the pilot scale, ensuring consistent production of high-quality UC-MSCs.

Conclusion

Our study provides valuable insights into the optimization of UC tissue processing protocols, the parameters for the enzymatic digestion method, and the comparative analysis of different isolation methods. We also demonstrated the stability of consecutive passaging using this method. Moreover, our scalable manufacturing processes enable large-scale production of high-quality UC-MSCs. These findings contribute to the advancement of UC-MSC-based therapies in regenerative medicine.

UC-MSCs

Separation method

Enzymatic digestion

Scale up

GMP

Stability

Quality control

Mesenchymal stem/stromal cells (MSCs) were first discovered by Friedenstein in bone marrow in the 1970s [1] and named by Caplan in 1991 [2]. Pittenger demonstrated that MSCs could differentiate into adipocytic, chondrocytic, or osteocytic lineages [3]. The International Society of Cellular Therapy (ISCT) proposed the minimal criteria for defining MSCs in 2006, and in 2015, they suggested incorporating immune functional assays into MSC potency release criteria [4] [5].

Thus far, a large number of clinical trials involving MSC therapy have been conducted worldwide, the number of which registered on Clinicaltrial.gov has exceeded 10,000 cases (www.clinicaltrial.gov). Additionally, approximately ten MSC-based cell therapy products have been approved for clinical use worldwide, such as Prochymal, Temcell, Alofisel, Cupistem, Stempeucel, and others [6]. Based on their immunomodulatory and tissue repair properties, MSCs have been used as a treatment for graft-versus-host disease (GvHD), systemic lupus erythematosus (SLE), inflammatory bowel disease (IBD), ischemic stroke (IS), Crohn’s disease (CD), knee osteoarthritis (KOA), spinal cord injury (SCI), and critical limb ischemia (CLI), among others [6–10].

To date, the predominant focus in clinical studies and product development has been on using MSCs derived from bone marrow (BM-MSCs) or adipose tissue (AT-MSCs). However, in recent years, significant attention has been directed toward UC-MSCs as a valuable source of mesenchymal stem cells. The availability of UC-MSCs is facilitated by their derivation from medical waste, typically discarded after birth, which ensures easy accessibility while minimizing pain and ethical concerns. Moreover, UC-MSCs possess noteworthy characteristics, such as low immunogenicity and high proliferation capability, thus enabling large-scale expansion. Importantly, long-term in vitro culture of UC-MSCs appears to have minimal impact on their phenotype and genetic stability. Clinical trials have not unveiled any long-term adverse effects or tumor formation associated with UC-MSCs. This further bolsters their potential for a broad range of clinical applications [11–17].

Due to the significant potential of UC-MSCs in regenerative medicine, extensive research has been conducted to optimize the manufacturing process [18, 19]. Efficient and reliable methods for UC-MSC isolation from the umbilical cord are critical to harness their therapeutic potential. Currently, the main methods commonly used are the explant method and enzymatic digestion method [20]. The explant method involves culturing small tissue fragments obtained from umbilical cord Wharton's jelly. This method offers the notable advantage of simplicity and minimizes external factors that could potentially damage the cells, thereby retaining the viability and functionality of UC-MSCs. However, this approach may require a longer culture time and typically yields a relatively lower number of cells than the enzymatic digestion method. On the other hand, the enzymatic digestion method utilizes specific enzymes to dissociate the matrix of umbilical cord tissues and release UC-MSCs. This technique provides a higher cell yield and requires a shorter culture time for expansion. Nevertheless, the enzymatic digestion process carries the risk of potentially damaging the cells if inappropriate digestive conditions are used, which may impact cell viability, phenotype or functionality [21–26]. Despite advancements, there is a lack of comprehensive studies focusing on developing good manufacturing practice (GMP)-based methods for UC-MSC separation and scaling up. This highlights the need for further research to establish standardized protocols that ensure the reproducibility and safety of UC-MSC manufacturing for clinical uses.

The objective of this study was to develop a GMP-compliant separation method for isolating Wharton's Jelly Derived Umbilical Cord MSCs. This includes the selection of GMP-compliant reagents, investigation of various process parameters, and conducting stability studies to ensure the consistent quality and safety of the isolated MSCs for therapeutic applications. Additionally, translational studies were conducted to bridge the gap between laboratory research and pilot-scale production. The ultimate goal is to establish a robust and standardized manufacturing process, ensuring that isolated MSCs can be utilized as safe and effective therapeutic agents.

UC tissue collection and preprocessing

Umbilical cord tissue was collected from Shenzhen Baoan District Maternity & Child Healthcare Hospital. The collection process was approved by the Medical Ethics Committee of Shenzhen Baoan District Maternity & Child Healthcare Hospital. The screening, collection, transportation, and infectious disease testing of parturients were carried out according to our standard operating procedures (SOPs). In short, the mothers were between the ages of 20–35 and free from infectious diseases and family genetic diseases and provided written informed consent. After collection, the umbilical cord (> 20 cm length) was transported to our manufacturing facility within 24 hours. Both umbilical cord blood and maternal blood samples are subjected to pathogen testing, which includes HBV, HCV, HTLV, TP, HIV, EBV, and CMV.

Then, the UCs were weighed, rinsed, decontaminated and divided into multiple segments. With the use of a surgical scalpel, the UC segments were opened to expose Wharton’s jelly and underlying blood vessels. Then, two arteries and one vein were removed, and Wharton’s jelly was extracted. Wharton’s jelly was rinsed again and minced into 1–4 mm³ fragments. These fragments can be further utilized to isolate MSCs using either the explant method or enzymatic digestion method. The explant method involves directly placing small tissue fragments onto the culture medium, while the enzymatic digestion method enzymatically breaks down the extracellular matrix to release desired cells for culture. The detailed process is described as follows.

Parameter selection of the enzymatic digestion method

To optimize the enzymatic digestion method for increased cell yield, various factors need to be taken into consideration, including the selection of enzymes, isolation parameters, and culture parameters. For application in clinical settings, the GMP grade enzyme Collagenase NB6 GMP (Nordmark Biochemicals) was recommended based on preliminary research conducted on various manufacturer brands. This optimized formulation includes collagenase class I and class II, as well as proteolytic activities such as neutral protease and clostripain. Isolating parameters primarily involve enzyme concentration, digestion time, temperature, and pH, while culture parameters focus on inoculation density and culture medium. It is worth noting that a temperature of 37°C and a pH range of 7.0-7.4 have been identified as optimal for enzyme activity. Consequently, a comprehensive assay design was developed, encompassing varying enzyme concentrations (0.2 PZ U/mL, 0.4 PZ U/mL, 0.6 PZ U/mL), digestion times (1 h, 2 h, 3 h), seeding densities (0.5 g tissue per flask, 1 g tissue per flask, 2 g tissue per flask), and culture mediums (MSC Serum- and Xeno-Free Medium (Biological Industries) + 2% hPL (Sexton Biotechnologies), 5% hPL, 10% hPL). Due to the complex composition of cell components immediately after digestion and significant variations among different samples, we selected postcultured passage 0 (P0) generation cells as the evaluation indicator for the results.

In addition to these factors, the enzymatic digestion method generally follows the previously described basic procedure. Briefly, the tissue fragments were transferred to a bottle, and twice the volume of collagenase was added. Then, the bottle was placed in a temperature-controlled shaking incubator for digestion, with digestion parameters set to 37°C and 150 rpm. After digestion, the mixture was neutralized by adding three times the volume of DPBS (Gibco). Filtration was then performed by passing the mixture through a 100-µm cell strainer (Corning). Subsequently, centrifugation was carried out at 1000 × g for 15 minutes to separate the suspended cells, the supernatant was removed, and the pellet was resuspended in DPBS. Another centrifugation step was performed at 400 × g for 10 minutes to further remove collagenase residue. After discarding the supernatant, the pellet was resuspended in culture medium. A sample was taken for cell counting (Vi-Cell Blu, Beckman), and the resulting cell suspension was transferred into a culture flask (Nunc) for incubation at 37°C in a humidified atmosphere with 5% CO₂. The first medium exchange was performed after 5 days, followed by subsequent medium exchanges every 3 days until days 10–15 or until the cell confluence reached 60%-80%. These MSCs were harvested using recombinant trypsin (Gibco) and designated P0. Cell morphology, quantity, viability, culture time, and immunophenotype were evaluated.

Comparative analysis of the explant and enzymatic digestion methods

A head-to-head comparative study was conducted to analyze the differences between the explant method and enzymatic digestion method. Briefly, 2 g UC tissue fragments were divided into two equal segments: one was directly added to culture medium and transferred to flasks (Nunc), and the other underwent the collagenase digestion process as described above. UC-MSCs were harvested at P0 and then seeded for continuous passage at a density of 5000 ± 10% cells per cm². Cell morphology, cell count at P0, culture duration, immunophenotype, and population doubling time of continuous passage were compared.

Stability of consecutive passaging UC-MSCs using the enzymatic digestion method

To assess the stability of the isolation method and establish the appropriate passage number for clinical application, P0 cells obtained using the enzymatic digestion method were consecutively passaged up to P9 at a density of 5000 ± 10% cells per cm². The evaluation included the change in cell morphology, population doubling time, cell viability and cell diameter with different passages. Cumulative population doublings were analyzed as well.

Manufacture and scaling up from laboratory scale to pilot scale

All the above experiments were conducted on a laboratory scale based on culture flasks (Nunc). To further evaluate whether the established method was suitable for GMP-compliant manufacturing, the production process was scaled up to a pilot scale based on a cell factory (Nunc), and UC-MSCs were passaged up to P5. A master cell bank (MCB) and working cell bank (WCB) were established at P1 and P3, respectively. The production and quality control of three batches was carried out in the GMP-compliant manufacturing facility of the Shenzhen manufacturing facility for personalized cell therapy of Beike.

In detail, first, a comparison was made between the monolayer cell factory and culture flask in terms of cell yield at P0. After enzymatic digestion of umbilical cord tissue fragments, the cells were cultured in both a 75 cm² culture flask and a 632 cm² monolayer cell factory at the same seeding density. Second, the P0 UC-MSCs were harvested using recombinant trypsin (Gibco) and seeded into another monolayer cell factory at a density of 5000 ± 10% cells/cm². After 3 days, when cell confluence reached 85%-95%, the cells were harvested and cryopreserved (7.5% DMSO, Wak-Chemie) to establish the master cell bank (P1). Third, the MCB cells were thawed and seeded into a monolayer cell factory at a density of 5000 ± 10% cells/cm², resulting in passage 2 (P2) cells. After a 3-day growth period, once cell confluence reached 85%-95%, the P2 cells were subsequently harvested and seeded into a 4-layer cell factory to obtain passage 3 (P3) cells, which were cryopreserved to establish the working cell bank. Finally, the WCB cells were thawed and seeded into a monolayer cell factory, resulting in P4 cells at a density of 5000 ± 10% cells/cm². After 3 days, when cell confluence reached 85%-95%, the P4 cells were then seeded into a 10-layer cell factory to obtain P5 cells, which were cryopreserved for further clinical use.

Quality control assays

Cell morphology

During the initial day of separating UC-derived MSCs, each culture medium exchange and cell harvest, cell observations were performed using an inverted microscope (IX73, Olympus). These observations encompassed cell morphology and cell confluence analysis.

Cell counting and viability

The cell numbers and viability were determined by an automatic cell counter (Vi-Cell Blu, Beckman) with the trypan blue exclusion method. After mixing Trypan Blue dye with the cell suspension and placing it into the cell counter, the counter captured 100 images for the analysis of cell count, viability, and cell diameter (µm).

Cell proliferation analysis

Cell proliferation analysis included calculating the population doubling time (PDT) and population doubling level (PDL). PDT was calculated by the formula X = T×log2/(logN-logX₀), where T is the time between initial plating and harvest for the respective passage, N is the total number of harvested cells, and X₀ is the total number of initial plating [27, 28]. PDL was calculated by the formula X=(logN-logX₀)/log2 [29].

Immunophenotype

Surface antigen phenotyping of UC-MSCs was assessed using flow cytometry (FACSCalibur™, BD Biosciences) and analyzed with CellQuest Pro software. The cells were stained with anti-human antibodies labeled with phycoerythrin (PE), allophycocyanin (APC), fluorescein isothiocyanate (FITC), or PerCP. The specific antibodies used for staining included CD73-PE, CD44-FITC, CD29-PE, CD166-PE, CD45-FITC, CD34-PE, CD14-FITC, CD79a-APC, HLA-DR-PerCP (BD Pharmingen), CD105-APC (eBioscience), CD90-FITC, CD31-PE, HLA-ABC-APC, CD80-FITC, CD40-PE, and CD86-APC (Biolegend). Prior to and following staining, samples were washed with 1× PBS. Isotype antibodies from the same manufacturers for mice or rats were used as controls.

Growth curve and cell cycle assay

UC-MSCs were initially seeded at a density of 10,000 cells per 12-well plate (Corning). Starting from Day 1 and continuing until Day 7, cells were harvested from three wells every day and counted using an automatic cell counter (Vi-Cell Blu, Beckman). The cell counts obtained during the logarithmic growth phase were utilized to calculate the population doubling time using the listed formula. Additionally, on Day 3, UC-MSCs were harvested specifically for the cell cycle assay. The Cycletest™ Plus DNA Kit (BD Biosciences) was employed to determine the cell cycle, and the assay was conducted according to the manufacturer's instructions. The proliferative index (PI) was calculated by the formula PI= (S + G2/M)/(G0/G1 + S + G2/M) [30].

CFU-F assay

UC-MSCs were initially seeded at a density of 100 cells per 6-well plate (Corning) in triplicate. After culturing for a period of 10–14 days, the cells were washed with 1× PBS, fixed in 100% methanol for 30 minutes, stained with 0.1% crystal violet for 45 minutes, and then rinsed in tap water 2–3 times. Aggregates consisting of 50 cells or more were defined as CFU-Fs (colony-forming units-fibroblasts). The data are reported as the total number of colonies per 100 cells [31–32].

Cellular senescence assay

The cellular senescence assay was performed using a senescence β-galactosidase staining kit (Beyotime) following the manufacturer’s instructions. Briefly, UC-MSCs were seeded in a 6-well plate (Corning) at a density of 2000–3000 cells and cultured for 72 hours. Subsequently, the cells were washed with 1× PBS, fixed at room temperature for 15 minutes, and then washed thrice with 1× PBS. Following this, the cells were incubated overnight at 37°C in a dry incubator with the X-gal staining mixture. After washing away the staining solution, the cells were visualized and captured using an inverted microscope (CKX53, Olympus). Cells exhibiting blue staining were identified as senescent cells. The percentage of senescent cells relative to the total number of cells represents the cellular senescence percentage.

Multilineage differentiation

The differentiation potential of UC-MSCs was assessed using the OriCell® osteogenic, adipogenic, and chondrogenic differentiation Kit (Cyagen Biosciences Inc.) according to the manufacturer's instructions. Briefly, for osteogenic and adipogenic differentiation, the cells were cultured in a 6-well plate at 37°C with 5% CO₂ until reaching approximately 70%-100% confluence. Subsequently, the culture medium was replaced with osteogenic or adipogenic differentiation medium and exchanged every 3 days. After 2 to 4 weeks of incubation, the cells were stained with Alizarin Red S solution or Oil Red O solution to evaluate osteogenic or adipogenic differentiation, respectively. For chondrogenic differentiation, (3–4) × 10⁵ cells were used to differentiate into chondrogenic cells for a period of 3 to 4 weeks, which were then stained with Alcian Blue solution.

The stained images were analyzed under an inverted microscope (100× magnification; CKX53, Olympus). In each differentiation assay, cells grown in regular medium were used as the negative control.

Karyotype analysis

Karyotyping was conducted using the Giemsa stain technique. First, cell division was halted in metaphase with 0.3 µg/mL colchicine (Solarbio) at 37°C for 2–3 hours. Afterward, the cells were washed and trypsinized, suspended in a warmed hypotonic solution (0.075 M KCl) and incubated for 30–40 minutes at 37°C. Following incubation, the cells were washed with a fixative solution consisting of a mixture of methanol and glacial acetic acid at a 3:1 ratio 3 times. Subsequently, the cells were resuspended in a fresh fixative solution and dropped onto clean slides, which were placed in ice water. To obtain G-bands, the slides were dried at a temperature of 70°C for a minimum of 3 hours. Next, the slides were immersed in a 0.01% trypsin solution for 3 minutes. They were then rinsed twice with saline solution before being stained using a 1:20 dilution of Giemsa solution (Bio Basic). Evaluation of the band quality was performed under a microscope with a magnification set at 100×. Mitoses were captured using specialized software, and a minimum of 20 metaphases were analyzed in each sample.

Mixed lymphocyte reaction

The mixed lymphocyte reaction assay was performed following the protocol described by K. Zhang et al. [33]. In brief, peripheral blood mononuclear cells (PBMCs) were thawed and suspended in PBS and labeled with carboxyfluorescein diacetate succinimidyl ester (CFSE). The CFSE-labeled PBMCs were stimulated with 10 ng/ml PHA and cocultured with hUC-MSCs (5:1) at 37°C and 5% CO₂ in a 12-well plate for 4–5 days. Following the incubation period, all lymphocytes were collected, and cell proliferation was determined by monitoring the gradual reduction in CFSE fluorescence using a BD FACS Calibur flow cytometer. PBMCs without labeling, PHA stimulation, or coculturing were used as negative controls.

Microbial safety test

Microbial safety testing was conducted following the guidelines set third in the Pharmacopoeia of the People's Republic of China (ChP). The bacterial and fungal tests employed a culture method, while the mycoplasma test utilized both the culture method and the indicated cell culture method. The endotoxin test was performed using the Limulus amebocyte lysate method.

Statistical analysis

Statistical analyses and graphs were performed using GraphPad Prism 8.0. Data are expressed as the mean ± standard deviation (SD). Significant differences between groups were assessed by paired t tests. A p value less than 0.05 was deemed to have statistical significance, while a p value less than 0.01 was considered highly statistically significant. Furthermore, a p value less than 0.001 was considered to indicate an extremely strong level of statistical significance.

Parameter selection of the enzymatic digestion method

The parameters studied in this research include enzyme concentrations (0.2 PZ U/mL, 0.4 PZ U/mL, 0.6 PZ U/mL), digestion times (1 h, 2 h, 3 h), seeding densities (0.5 g tissue per flask, 1 g tissue per flask, 2 g tissue per flask), and culture mediums (2% hPL, 5% hPL, 10% hPL). UCs were collected from a total of 9 donors; 3 donors’ UCs were utilized for studying enzyme concentrations and digestion times, another 3 were used for investigating digestion times, and the remaining 3 were used for culture medium studies.

An orthogonal experimental design was employed to investigate the effects of enzyme concentrations and digestion times (Fig. 1A). After digestion with collagenase NB6 GMP and culture for 10–15 days, P0 UC-MSCs were harvested. The mean quantity of P0 UC-MSCs among the nine groups in the three samples was 1.83×10⁶ ± 1.30×10⁶, 5.86×10⁵ ± 5.39×10⁵, and 6.66×10⁵ ± 8.78×10⁵, indicating inherent differences among the samples. To facilitate better comparison, we transformed the cell quantities into scores. A group of each sample was assigned a maximum score of 100 based on the highest cell quantity observed. The remaining groups were then proportionately adjusted to derive their respective scores in each group. The average cell score was calculated for each parameter, as illustrated in Fig. 1B-C. Concentrations at 0.4 PZ U/mL led to an observed increase in cell quantity, while concentrations at 0.6 PZ U/mL showed a decreasing trend. Similarly, digestion times of 3 hours were associated with an increase in cell quantity, whereas digestion times of 4 hours exhibited a declining trend. The expression of surface markers on the P0 UC-MSCs obtained through the enzyme digestion method was found to be consistently stable across different samples and groups (Fig. 1D).

In addition to the digestion parameters, the culture parameters also influence the quantity of P0 UC-MSCs. After digesting 0.5, 1, and 2 g of Wharton’s jelly, primary cells were obtained in amounts of 4.05×10⁴ ± 5.89×10³, 1.05×10⁵ ± 1.40×10⁴, and 2.32×10⁵ ± 9.03×10⁴, respectively. Subsequently, the quantities of P0 cells generated were 2.14×10⁶ ± 1.82×10⁶, 3.64×10⁶ ± 2.84×10⁶, and 1.15×10⁷ ± 7.22×10⁶, respectively. A clear linear correlation was observed between the grams of Wharton’s jelly used and the primary cells, as indicated by the high coefficient of determination (R² = 0.97). Correspondingly, the quantity of P0 UC-MSCs generated also exhibited a linear relationship with the grams of the initial Wharton’s jelly used (R² = 0.88) (Fig. 1E-F). However, no correlation was found between the seeding density of primary cells and the quantity of P0 UC-MSCs among the different samples, as shown in Fig. 1G.

Interestingly, it was observed that the addition of 2% hPL had an equivalent effect to that of 5% hPL. The quantity of P0 UC-MSCs obtained when 2% hPL was added was 1.31×10⁷ ± 2.25×10⁶, while it was 1.32×10⁷ ± 2.89×10⁶ for the 5% hPL group. In contrast, increasing the hPL concentration further to 10% resulted in a decrease in the quantity, with the cell count reaching 1.11×10⁷ ± 1.86×10⁶ (Fig. 1H). This intriguing result suggests that there may be an most suitable concentration of hPL for promoting the expansion of UC-MSCs, beyond which higher concentrations may have a negative impact on cell proliferation.

Comparative Analysis of the Explant and Enzymatic Digestion Methods

Three UCs were collected, and the Wharton’s jelly obtained from each sample was separated into two groups. One group was used for collagenase digestion culture, while the other group was used for explant culture. The seeding density for both methods was 1 gram per flask. The enzymatic digestion method initiated cell proliferation between days 6–8 and exhibited uniform growth across the entire culture surface, while the explant method initiated cell proliferation between days 8–10 and demonstrated high-density growth in the center of the tissue, extending outward. It is noteworthy that exfoliated cells were observed prior to cell collection in the explant method culture. Additionally, both types of P0 UC-MSCs displayed a spindle-shaped morphology (Fig. 2A).

The average culture duration for P0 cells obtained through the enzymatic digestion method was 11.37 ± 1.21 days, which was highly significantly shorter than the culture duration of 13.66 ± 1.06 days obtained through the explant method (p < 0.01). The quantity of P0 UC-MSCs harvested through the enzymatic digestion method was 3.53×10⁶ ± 9.02×10⁵ cells per gram of Wharton’s jelly, which was slightly lower than the quantity of P0 UC-MSCs obtained through the explant method (4.01×10⁶ ± 1.06×10⁵ cells per gram of Wharton’s jelly). Therefore, the enzymatic digestion method demonstrated faster cell proliferation (Fig. 2B-C).

No significant differences were observed in the average PDT during passaging from P1 to P4 or in the viability and immunophenotype of P0 and P4 UC-MSCs between the two different methods (Fig. 2D, E, F). The PDT values ranged from 17.80 to 22.31 hours among different samples, and the viability of P0 and P4 UC-MSCs was above 85%. The expression levels of positive markers were consistently above 95%, while the negative markers were consistently below 2% for both. These outcomes collectively demonstrate that both methods yield UC-MSCs of excellent quality and consistency.

Stability of consecutive passaging UC-MSCs using the enzymatic digestion method

To evaluate the stability of UC-MSCs using the enzymatic digestion method, a total of five umbilical cords (UCs) were collected, and the cells were continuously passaged up to P9. The PDT of the cells remained within the range of 15–30 hours, gradually decreasing from P0 to P3, with P3 showing the shortest doubling time (16.92 ± 1.26 h) and the fastest cell proliferation. However, starting from P6, the doubling time gradually increased, indicating a slowdown in cell expansion beyond the sixth passage (Fig. 3A).

The cell viability among different passages remained relatively consistent with minor variations, consistently exceeding 85%. Notably, the cell viability was significantly higher in P2 (98.82% ± 1.70%) than in the other groups, showing best cell activity during P2 (Fig. 3B).

The cell diameter ranged from 14.58 to 17.60 µm among different passages. From P5, the cell diameter gradually increased. Each passage from P0 to P4 cells showed significant differences compared to P6, as did each passage from P0 to P6 when compared to each passage from P7 to P9 (Fig. 3C).

With each passage, there was a consistent and incremental increase in cumulative population doublings (CPDs), demonstrating continuous and reliable cell proliferation. The CPDs were recorded as 20.42 ± 1.05 at P5 and further increased to 36.66 ± 1.99 upon reaching P9 (Fig. 3D). The average population doublings (PDs) across all passages were 4.07 ± 0.54.

Although all cell passages exhibited a spindle-shaped morphology, notable differences were observed. P1 displayed a similar morphology to P5, with cells appearing small, short, and thin. In contrast, P9 cells appeared larger, longer, and more voluminous (Fig. 3E).

These findings underscore the remarkable ability of UC-MSCs to consistently expand and proliferate over successive passages. Notably, P2 to P5 demonstrated superior cell activity, highlighting their potential clinical utility.

Manufacture and scaling up from laboratory scale to pilot scale

The scale-up study from culture flasks to cell factories was conducted on three additional UC samples. The UC sample information is shown in Fig. 4A. UC-MSCs were isolated using the enzymatic digestion method. After being cultured for 10–15 days, the quantity of UC-MSCs (P0) obtained from each sample using the monolayer cell factory was determined to be 3.78 ×10⁷, 2.05 ×10⁷, and 9.57 ×10⁷ cells, respectively. In comparison, the quantity of P0 UC-MSCs obtained from the ^{25 cm2} flask was 3.75 ×10⁶, 2.75 ×10⁶, and 1.06 ×10⁷ cells, respectively. This indicates that the quantity of P0 cells per cm² in the flask was two to three times higher than that in the cell factory, clearly demonstrating that the culture vessel surface area has an impact on cell quantity (Fig. 4B).

In Fig. 4C, the mean PDT, CPD, and proliferation fold of all passages cultured using the cell factory are presented. Notably, UC3 showed remarkable proliferation ability, with an average PDT of 18.24 ± 3.83 hours across all passages and a proliferation fold reaching 936,780 from P0 to P5. As shown in Fig. 4D, the estimated average quantity of UC-MSCs was projected to be 1.07 ×10⁹ at P1, 2.03 ×10¹¹ at P3, and 3.23 ×10¹³ at P5. Furthermore, the estimated average quantity of UC-MSCs per gram when reaching P5 was 9.51 ×10¹¹.

Quality control (QC) assessments were conducted on P1 (MCB), P3 (WCB), and P5 (DP), encompassing a range of evaluations, such as cell viability, immunophenotype, growth curve, cell cycle assay, CFU-F assay, cellular senescence assay, karyotype analysis, multilineage differentiation potential, mixed lymphocyte reaction, and microbial safety testing. The viability was 93.64 ± 6.83% for P1, 89.70 ± 2.95% for P3 and 87.32 ± 0.81% for P5, with all values above 85% (Fig. 5A). The expression levels of CD73, CD90, CD105, CD44, CD29, and CD166 markers were uniformly greater than 95%, indicating positive immunophenotype characteristics. Conversely, the expression of CD45, CD34, CD79a, CD14, CD31, and HLA-DR markers was less than 2%, demonstrating the minimal presence of non-MSC contaminants. HLA-ABC exhibited positive expression, with P5 samples showing expression levels exceeding 95%, while P1 and P3 displayed expression levels above 60%. The costimulatory molecules CD80, CD40, and CD86 exhibited low expression levels below 5%, with CD80 and CD40 showing minimal expression and CD86 being relatively higher than CD80 and CD40. The expression of the therapeutic function-related marker CD146 was moderate and varied among the three samples, showing a decrease with an increase in passages (Fig. 5B-C).

To further evaluate the activity and growth potential of UC-MSCs, several assays were conducted, including growth curve, cell cycle, CFU-F, and cellular senescence assays. The mean PDT calculated from the growth curve was 28.42 ± 1.46 hours for P1, 26.70 ± 1.87 hours for P3, and 28.98 ± 3.45 hours for P5 (Fig. 5D-E). The proliferative index (PI) calculated from the cell cycle assay indicated percentages of 48.51 ± 4.62% for P1, 48.34 ± 0.34% for P3, and 41.66 ± 5.16% for P5 (Fig. 5F-G). Additionally, the CFU-F assay revealed the formation of 21.78 ± 4.84 colonies for P1, 16.67 ± 7.51 colonies for P3, and 17.67 ± 7.57 colonies for P5 (Fig. 5H). Furthermore, the percentage of cells showing β-galactosidase activity, an indicator of cell senescence, was found to be 0.23 ± 0.19% for P1, 0.14 ± 0.11% for P3, and 0.56 ± 0.26% for P5 (Fig. 5I). These results indicated overall favorable cell activity and growth potential in our study, albeit with a gradual decline observed as passages increased.

Karyotype analysis was performed at P5 to assess the genetic stability of UC-MSCs. The results showed no numerical or structural chromosome abnormalities (Fig. 6A). In addition, microbial safety tests conducted on P1, P3, and P5 samples, including evaluations for bacteria, fungi, mycoplasma, and endotoxin, all yielded negative results (Fig. 6B), providing assurance regarding the safety of our facility.

In addition to the aforementioned analyses, a biologic activity assay was conducted in this study. The P5 UC-MSCs exhibited multilineage differentiation potential, successfully differentiating into osteogenic, adipogenic, and chondrogenic lineages (Fig. 6C). In addition, a mixed lymphocyte reaction (MLR) assay was performed to evaluate the inhibitory effect of UC-MSCs on lymphocyte proliferation. The results demonstrated a favorable ability of P1, P3, and P5 UC-MSCs to inhibit lymphocyte proliferation, with percentages of 66.78 ± 6.79%, 72.72 ± 5.95%, and 70.29 ± 5.57%, respectively. Notably, no significant differences were observed between different passages, indicating that there was no decrease observed with an increase in the number of passages (Fig. 6D-E).

These observations highlight the successful scale-up study, demonstrating a seamless transition from culture flasks to cell factories. This transition allows for the large-scale production of UC-MSCs, making them suitable for a wide range of clinical applications.

Currently, UC-MSCs have been extensively utilized in clinical trials [11, 12, 34]. However, there has been limited focus on developing SOPs for GMP to ensure consistent quality of these cells. In this study, we established a standardized method for isolating UC-MSCs, which is outlined as follows: First, Wharton's jelly was extracted and cut into tissue fragments measuring 1–4 cubic millimeters. GMP-grade digestive enzyme (collagenase NB6 GMP) was added at a concentration of 0.4 PZ U/mL, and the digestion process was conducted for a duration of 3 hours. After digestion, the cell suspension was diluted with PBS and filtered using a cell strainer. For Wharton's jelly weighing above 20 g, the cells obtained from digestion could reach a minimum of 2×10⁶ cells to be seeded in a single-layer cell factory. Subsequently, the culture medium was exchanged after 5 days of culture, with subsequent medium exchanges occurring every 3 days until days 10–15. At this stage, P0 cells can be harvested for further production. The parameters within this method were optimized to ensure the generation of high-quality UC-MSCs.

The explant method and enzymatic digestion method are the two main approaches for isolating UC-MSCs [20]. In this work, we conducted a parallel comparison using the same umbilical cord and the same weight of Wharton’s jelly. Our findings revealed that the enzymatic digestion method demonstrated a faster start-up and a shorter culture time during the initial passage (P0). However, after subsequent passages, there were no significant differences between the two methods in terms of cell proliferation, cell viability, and immunophenotype. This suggests that damage caused by enzymatic digestion can be recovered during the culture process. Additionally, the enzymatic digestion method can be easily translated into an automation platform, making it more suitable for GMP-compliant processes in a standardized manufacturing setting [35, 36].

Our findings indicated a positive correlation between the initial cell seeding density and the number of cells collected at P0 within the same sample using the enzymatic digestion method. However, we observed no correlation across different samples due to the individual differences and intricate composition of umbilical cord cells, posing a challenge in determining an appropriate initial cell seeding density. The UC tissue comprises two umbilical arteries, one umbilical vein, and the allantois embedded in Wharton’s jelly and is surrounded by a single layer of amnion. Multiple cell types are present within the UC, including mesenchymal stem cells (MSCs), endothelial cells, epithelial cells, fibroblasts, and cord blood cells. MSCs in Wharton’s jelly are primarily found in the perivascular region nearest to the umbilical vessels, with fewer in the intervascular region and the least in the subamniotic region [37, 38]. Research studies have shown that the colony-forming unit (CFU) derived from primary human umbilical cord perivascular (HUCPV) cells was 1 CFU for every 333 cells [39], while another study reported a CFU-F frequency of 1:1609 in nucleated cells from the umbilical cord [40]. These discoveries further support our research results.

Human platelet lysate (hPL) is widely used as an alternative to fetal bovine serum (FBS) in the ex vivo expansion of MSCs. hPL contains a high concentration of growth factors, adhesion molecules, and chemokines, making it suitable for the production of GMP-compliant cell products [41–44]. Compared to FBS, hPL offers several significant advantages, including enhanced proliferation behavior, reduced population doubling time, preservation of clonogenicity, increased CFU-F size, maintenance of characteristic immunophenotype, preserved in vitro trilineage differentiation capacity, maintained in vitro T-cell immunosuppression, and absence of in vivo tumorigenicity. More importantly, the use of hPL eliminates the risks associated with the transmission of animal-derived viruses [45].

In our study comparing the effects of different concentrations (2%, 5%, and 10%) of hPL on primary cell expansion, interesting outcomes were observed. The results demonstrated that both 2% and 5% concentrations showed similar levels of cell expansion. However, using a 10% concentration resulted in decreased cell expansion. This finding is consistent with previous studies. According to Shansky et al., a 5% concentration of hPL was found to be more effective than both 1% and 10% concentrations in supporting AT-MSC growth. Similarly, Azouna et al. reported that the PDT of 5% hPL was not significantly lower than that of 10% HPL or a combination of 10% FBS and 5% hPL [46, 47]. A meta-analysis has shown that 5% hPL is superior to 10% FBS [48], and Kirsch et al. found that even a lower concentration of 2.5% hPL exhibited a higher proliferation and differentiation rate compared to 10% human serum (HS) or 10% fetal calf serum (FCS) in AT-MSCs [49]. Notably, the discussed above studies conducted their experiments by supplementing basal media, such as α-MEM or DMEM, with additional components. Moreover, it was discovered that when using defined serum-free mesenchymal stem cell media that have been optimized for growth factors, the concentration of hPL can be further reduced to 1% or even as low as 0.5% [50–51]. This observation may provide an explanation for the findings of our study, suggesting that the efficacy of hPL in supporting MSC growth and function can be maximized even at lower concentrations when used in conjunction with serum-free media formulations.

According to a study by Ikebe et al., the use of MSCs in clinical trials from 2007 to 2013 showed that 23% of trials used cells from passage 1 or less, 71% used cells from passages 1–5, and only 6% used cells from passages over 5 [52]. Sareen et al. demonstrated that an increase in passage number (from P3 to P7) in cell culture did not have a significant effect on the immune privilege of MSCs [53]. However, another study found that cells gradually lost their typical fibroblast-like spindle shape from P3 to P8, resulting in elevated morphological abnormalities and inhomogeneity. The cell population doubling rate also decreased [54]. Zhao et al. found that cells at P3, P6, and P15 showed similar morphology, biomarker expression, and cytokine secretion. However, the therapeutic effect on aGVHD in vivo declined at P15 [55]. Yu et al. observed that MSCs grew well for 20 population doublings (PD) but experienced cellular senescence at approximately 40 PD [56]. Based on our stability study, it was found that passages 2 to 5, with a PD of less than approximately 20, were the better passages in terms of high viability and proliferation ability, particularly passage 2 or 3. For allogeneic therapy requiring an abundant number of cells, passages 4 or 5 may be the most suitable.

When transitioning the culture of MSCs from flasks to cell factories, whether at P0 or subsequent passages, the proliferation rate tends to decrease. This can be attributed to the heterogeneity of the physical and chemical environment, as well as the emergence of concentration gradients in cell factories, mainly due to gas exchange occurring at the medium/headspace gas interface [57].

However, despite the decline in the cell proliferation rate, successful scale-up of MSC manufacturing has been achieved in our study, resulting in high-quality drug products. These included high cell viability, maintenance of a stable immunophenotype, low cellular senescence percentage, stable karyotype, and maintenance of multilineage differentiation potential. Furthermore, MSCs showed potent inhibition of lymphocyte proliferation and were free from microbial contamination. The theoretical quantity of cell formulation reached 10¹³ from one UC.

Cell surface markers are one of the key indicators used to identify MSCs. In addition to testing the marker expression defined by the International Society for Cellular Therapy (ISCT) [4], we also detected the expression of other markers, such as adhesion molecules CD44, CD29, and CD166, functional marker CD146, immunogenic markers HLA-ABC, and costimulatory molecules CD40, CD80, and CD86. Our results revealed that CD73, CD90, CD105, CD44, CD29, and CD166 were expressed at levels higher than 95%, which are currently used to define MSCs [58]. On the other hand, CD45, CD34, CD79a, CD14, and CD31 were expressed at levels lower than 2%, implying a high MSC purity. Furthermore, we found that HLA-DR and costimulatory molecules CD40, CD80, and CD86 were expressed at low levels, while HLA-ABC was expressed positively. HLA-ABC plays a role in protecting MSCs from destruction by natural killer cells, while MHC-II helps in evading immune recognition by T cells. The costimulatory molecules CD40, CD80, and CD86 are part of the second signaling system for T lymphocyte activation. These findings indicate that UC-MSCs manufactured by our method are unlikely to trigger an immune response and can evade host immune attack in vivo [59, 60]. Recent research has shown that CD146 is a potency marker. The CD146⁺ subpopulation has enhanced immunosuppressive capacity, resulting in improved therapeutic outcomes [61, 62]. We found that the expression of CD146 gradually decreased with increasing passages, indicating that higher passages should be unsuitable for clinical practices. Interestingly, the expression of all the mentioned above markers aligns with the results of Miryam Mebarki et al. [12].

It is worth noting that although our study established GMP-compliant separation methods, manual operations involved in extracting Wharton's jelly and mincing the umbilical cord increase the risk of contamination. Some successful research studies have utilized the entire umbilical cord for digestion without the need to open and remove the vessels to obtain UC-MSCs [63, 64]. However, it is important to mention that these studies were conducted using umbilical cords of approximately 1 cm in length, and the results for longer lengths are still unknown. Therefore, further optimization of the separation method is still required to automate the process and improve consistency. Additionally, our study highlights certain limitations associated with the use of cell factories, suggesting a shift toward the use of bioreactors. Bioreactors allow for precise monitoring and tight regulation of essential culture parameters, including pH value, temperature, dissolved O₂, and CO₂ levels [57, 65]. This trend of utilizing bioreactors offers promising advantages in terms of enhanced control and scalability for MSC production.

In summary, our study successfully established a GMP-compliant separation and culture method for isolating Wharton's jelly derived MSCs from the umbilical cord. We have extensively optimized the parameters and conducted scaled-up manufacturing. The resulting cell product underwent thorough evaluations for identity, purity, viability, potency, proliferative capacity, genomic stability, and microbiological safety. The establishment of this method represents a significant advancement in the field and holds great promise for clinical translation.

UC-MSCs

Umbilical cord Wharton's jelly derived mesenchymal stem cells

GMP

Good manufacturing practice

hPL

human platelet lysate

ISCT

International Society of Cellular Therapy

BM-MSCs

Bone marrow mesenchymal stem cells

AT-MSCs

Adipose tissue mesenchymal stem cells

SOPs

Standard operating procedures

MCB

Master cell bank

WCB

Working cell bank

PDT

Population doubling time

PDL

Population doubling level

Proliferative Index

CFU-Fs

Colony-forming units-fibroblasts

PBMCs

Peripheral blood mononuclear cells

CFSE

Carboxyfluorescein diacetate succinimidyl ester

CPDs

Cumulative population doublings

PDs Population doublings

MLR

Mixed lymphocyte reaction

FBS

Fetal bovine serum

Human serum

FCS

Fetal calf serum.

Acknowledgments

The authors would like to extend their heartfelt thanks to the Shenzhen Baoan District Maternity & Child Healthcare Hospital for their valuable support and cooperation in facilitating the collection of umbilical cord samples.

Authors’ contributions

CWL designed the study and wrote the manuscript. FTH, XPZ, QFW, and FZ carried out the isolation and manufacturing of UC-MSCs. GYS and JFW performed the quality control assays. All the authors have read and approved the final manuscript.

Funding

The research was supported by Beike Biotechnology Co., Ltd. The funding body played no role in the design of the study and collection, analysis, and interpretation of data and in writing the manuscript.

Availability of data and materials

All data generated during this research are included in this published article.

Ethics approval and consent to participate

All UCs were obtained from healthy donors after written informed consent was obtained. The collection process was approved by the Medical Ethics Committee of Shenzhen Baoan District Maternity & Child Healthcare Hospital. The research project titled “Process Study and Animal Experiment of Intravenous Umbilical Cord Mesenchymal Stem Cell Therapy for Refractory Systemic Lupus Erythematosus” was approved on January 12, 2023 (Approval number: LLSCHY-2022-04-01-07).

Consent for publication

Not applicable

Competing interests

The authors declare that they have no competing interests.

Author details

¹Shenzhen Beike Biotechnology Co., Ltd., Shenzhen, Guangdong 518000, People’s Republic of China. ²National Engineering Research Center of Foundational Technologies for CGT Industry, Shenzhen, Guangdong 518000, People’s Republic of China. ³Shenzhen Kenuo Medical laboratory, Shenzhen, Guangdong 518000, People’s Republic of China.

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Development of a GMP-Compliant Separation Method for Isolating Wharton's Jelly Derived Mesenchymal Stromal Cells from the Umbilical Cord

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Journal Publication

Version 1

Abstract

Background

Methods

Results

Conclusion

Figures

Introduction

Materials and Methods

UC tissue collection and preprocessing

Parameter selection of the enzymatic digestion method

Comparative analysis of the explant and enzymatic digestion methods

Stability of consecutive passaging UC-MSCs using the enzymatic digestion method

Manufacture and scaling up from laboratory scale to pilot scale

Quality control assays

Cell morphology

Cell counting and viability

Cell proliferation analysis

Immunophenotype

Growth curve and cell cycle assay

CFU-F assay

Cellular senescence assay

Multilineage differentiation

Karyotype analysis

Mixed lymphocyte reaction

Microbial safety test

Statistical analysis

Results

Parameter selection of the enzymatic digestion method

Comparative Analysis of the Explant and Enzymatic Digestion Methods

Stability of consecutive passaging UC-MSCs using the enzymatic digestion method

Manufacture and scaling up from laboratory scale to pilot scale

Discussion

Conclusion

Abbreviations

Declarations

References

Status:

Journal Publication

Version 1