Cell Isolation

Introduction

At the foundation of any cell therapy development and manufacturing workflow, the quality of the starting cellular material directly impacts the final product’s viability and the efficacy of patient treatment. Most cell isolation methods, specifically peripheral blood mononuclear cell (PBMC) isolations, are currently performed using open systems, which can contribute to errors and contamination, resulting in the failure to produce a viable cell therapy. Additionally, donor product variability can lead to differences in cell composition, cell viability, and sensitivity during the expansion stages of the manufacturing process. For a multitude of reasons, not all patients, including healthy donors, are able to effectively mobilize functional T-cells at the desired numbers suitable for the process. Therefore, T-cell isolation workflows must be flexible to allow for various modifications, while still yielding a standardized CAR T-cell product, regardless of the input material harvest from the patient or donor. The ideal isolation workflow would also be automated, closed, and consistent.

This section will discuss the isolation of PBMCs and selection and activation of the T cell populations (CD3⁺) to be engineered. This section will also discuss alternative cell approaches using induced pluripotent stem cell (iPSC)-derived cells and natural killer (NK) cells, which are potential solutions to overcome issues with the more traditional approach.

Healthy donor characteristics

The development of an allogeneic cell therapy begins with the isolation of T-cells from donated blood in a process known as leukapheresis. To go to a clinical trial, a sufficient amount of donated blood is needed with the correct cell composition and phenotypes. For an allogeneic therapy, the ideal donor blood is preferably from a young individual. The donated blood composition should have a low percentage of monocytes and neutrophils (specifically granulocytes), and cells should demonstrate an efficient doubling time. The donor blood should have an immunological profile consisting of a normal CD3⁺ cell number, a balanced CD4/CD8 ratio, a sufficient number of stem memory cells, and expression of CD62L⁺CCR7⁺T-cells.

As expected, cell therapy developers and manufacturers want the highest purity of T cell population possible and sometimes look for specific subsets of T-cells. Poor quality starting material may result in an inability to use it for cell therapy processing and ultimately, failure of the donor product reaching the clinic. However, efficient cell isolation methods that are reliable with high yields and purity can alleviate the pressure on the quality of donor blood.

PBMC isolation

Once donated blood is obtained and characterized, the next step is to isolate PBMCs. Typically, the isolation methods are characterized as either open or closed, depending on the amount of direct user interaction needed. Closed methods are preferred because of a decreased risk for contamination and user error, and for some clinical applications, might be required.

The most well-known practice to isolate PBMCs relies on density gradient centrifugation using Ficoll medium [1]. Although this method successfully isolates PBMCs from red blood cells, the isolated PBMCs retain contaminants such as granulocytes, monocytes, and even some residual red blood cells. Most density gradient centrifugation isolations are performed using an open system, which makes the procedure prone to errors, contamination, and user-to-user variability. While automated closed systems for density gradient centrifugations exist, these systems tend to lose cells, lowering yields due to the lack of system flexibility and can be significantly more expensive.

A more recent closed system approach relies on counterflow centrifugation, which separates cells based on their size and density. Systems using this technology (e.g., Gibco CTS Rotea system) suspend cells in a fluidized bed by exerting a constant flow force against centrifugal forces (Figure 1). The suspended cells are gently concentrated without forming a pellet and then washed with very high recoveries. Using elutriation, dead cells can be removed to optimize viability of the population. Adjustments to centrifugal speed and flow rate allow for cells to be fractionated based on size and density, with minimal shear.

The Rotea counterflow centrifugation instrument is capable of separating cells through a process called elutriation, washing them, and concentrating a subpopulation of cells to the user’s specifications.

Figure 1. Principle behind counterflow centrifugation (CFC). The closed system CTS Rotea instrument relies on counterflow technology to separate cells based on size and density. A. Cell loading: With “balanced” g-force and counterflow parameters, input material comprising media and cells is introduced to the CFC chamber via the central tube. B. Elutriation: Larger/denser cells are captured in the fluidized bed, while smaller/less dense cells and debris pass through and are “eluted” through the top of the CFC chamber. C. Media exchange/washing: Wash buffer is pumped through the fluidized bed, replacing the original media in the input product. Note: The fluidized bed enables very fast and efficient washing. D. Cell concentration: Washed and concentrated cells are now recovered from the CFC chamber by simply reversing the pump and extracting the concentrate via the internal tube. More details and video on counterflow centrifugation technology can be found here.

Density gradient centrifugation and counterflow centrifugation produce comparable results (Figure 2). However, distinct advantages of the counterflow centrifugation methodology are that it can be performed more quickly, and most importantly, in the more desirable closed system setting. Table 1 summarizes the PBMC isolation methods.

Flow cytometric analysis of cells separated using a traditional density gradient method compared to cells separated using the CTS Rotea system shows that the Rotea system is capable of separating distinct cell types from a mixed population.

Click image to enlarge

Figure 2. Closed counterflow centrifugation system versus open density gradient system for isolation of PBMCs from red blood cells. A single-donor leukopak was divided into two, and PBMCs were separated using (A) density gradient centrifugation using Ficoll polymer or (B) counterflow centrifugation using the CTS Rotea system. The CTS Rotea system can isolate PBMCs from a leukopak in less than 30 minutes, with equivalent performance to the density gradient system and the added benefit of closed processing.

Table 1. Comparison of PBMC isolation methods.

	Density gradient centrifugation	Counterflow centrifugation
Advantages	Well-known Widely used Inexpensive (open system only)	Shortened processing times Protocol flexibility Less user error and variability Less contamination Automation-capable
Disadvantages	Long and tedious process More expensive (closed system)	More expensive (closed system)

T-cell isolation and activation

Following PBMC isolation, the next step in the process is the isolation of T-cells to remove any lingering contamination of other cell types and to improve product specificity. This is then followed by activation. The inherent variability with allogeneic donor blood leads to an inability to differentiate between cell types. To overcome this problem, magnetic bead-based approaches for selection of T-cell populations have been developed. These approaches use magnetic beads conjugated to antibodies that recognize T cell surface markers and bind to them. When placed in close proximity to a magnet, the bead-T-cell complex binds and is held while unwanted cell types can then be washed away. Once isolated, the T-cells can be released from the beads (see Figure 3).

Several magnetic bead products are commercially available. Some of these platforms can be used for very specific cell populations (e.g., CD4/CD8⁺ or CD62L⁺), which are activated later through other means. Another system, Gibco CTS Dynabeads CD3/CD28, provides both the primary and co-stimulatory signals required for activation and expansion of T-cells, eliminating the need for a separate activation step and reducing the potential to introduce contamination.

CTS CD3/CD28 Dynabeads magnetic beads, when used with a magnet, help scientists isolate cells that express CD3/CD28+ markers from other cells that do not express those markers, by binding to those cells and allowing the non-bound cells to be washed away.

Figure 3. Use of CTS Dynabeads CD3/CD28 to isolate and activate T cells for subsequent engineering. PBMCs are harvested and are activated using the CTS Dynabeads CD3/CD28 (bead:cell ratio of 3:1) in cell culture bags. The cells and beads in culture media are incubated for 30 minutes on a rocker. After the 30 minutes, the culture bag is placed on the CTS DynaMag Magnet and CD3/CD28⁺ cells attached to beads are captured. The supernatant and non-specified cells are discarded and fresh media with cytokines is added for simultaneous activation and expansion of the T-cells. Using this approach, CD3⁺CD28⁺ T-cells are isolated with over 90% recovery, uniformly stimulated (>95% CD25⁺), and highly pure population with over 95% CD3⁺, with no need for antigen presenting cells (APCs). For more details on this process, see One-step isolation and activation of naive and early memory T cells with CTS Dynabeads CD3/CD28

iPSC-derived CAR NK cells

One major issue with allogeneic workflows is insufficient cell numbers or starting material to create a product for patient infusion. To overcome this limitation, a revolutionary approach has been developed to derive natural killer (NK) cells or T cells from induced pluripotent stem cells (iPSCs). The use of NK cells reduces rejection barriers (e.g., graft-vs-host disease) and NK cells can be generated from several different sources such as umbilical cord blood, bone-marrow, human embryonic stem cells, and iPSCs.

A significant advantage of NK cells is that unlike T cells, they exhibit improved survival after killing multiple target cells. NK cells also produce a different profile of cytokines than T cells. The cytokines produced by T-cells cause cytokine release syndrome (CRS), a life-threatening condition that has been observed in some patients using an adoptive cell therapy, and this can potentially be avoided with the use of NK cells.

The most common cause of allogeneic therapy rejection is due to differences in the human leukocyte antigen (HLA) class I gene of iPSC-derived products, resulting in a mismatch between the donor and recipient. T-cells regularly interact with HLA complexes, thus any changes to the HLA complex on the surface would be an indication that the substance is foreign. Conversely, NK cells can exhibit cytotoxicity toward different tumor targets in an HLA-independent manner which could mitigate this mismatch between donor and recipient. The isolation of primary NK cells is difficult and complex and can lead to low yields, making iPSC-derived CAR NK cells a more attractive choice for allogeneic workflows. To this end, the main goal is to identify an iPSC line(s) that can avoid allogeneic rejection and reduce the time to move cell therapies into the clinic.

Summary

Preparation of allogeneic CAR T-cells requires identification of healthy third-party donors and isolation of a sufficient number of T cells for the subsequent engineering steps. Ideally, isolation would be performed in a closed system environment that can provide flexibility for modifications to easily account for differences in source material. Recent advancements have investigated use of other sources or cell types (i.e., iSPC-derived NK cells) to further expand and improve the use of the CAR T-cell technology.