Method Validation for Automated Isolation of Viable Peripheral Blood Mononuclear Cells

Abstract

Background:

This article is part of a series of publications providing formal method validation for biospecimen processing in the context of accreditation in laboratories and biobanks. We report the optimization and validation for fitness-for-purpose of automated and manual protocols for isolating peripheral blood mononuclear cells (PBMCs) from whole blood, and compare the two methods.

Methods:

The manual method was optimized for whole blood centrifugation speed, gradient type (Ficoll, Leucosep, CPT), and freezing method (Mr Frosty, Controlled Rate Freezing). Various parameters of the automated protocol using a CPT gradient on a Tecan liquid handler were optimized. Optimal protocols were validated in parallel for reproducibility and robustness. Optimization and validation were assessed in terms of cell yield, viability, recovery, white blood cell (WBC) subpopulation distribution, gene expression, and lymphoblastoid cell line (LCL) transformation.

Results:

An initial centrifugation of whole blood at 2000 g was considered optimal for further processing, allowing isolation of plasma and PBMCs from a single sample. The three gradients gave similar outcomes in terms of cell yield, viability, and WBC subpopulation distribution. Ficoll showed some advantages and was selected for further evaluations. Optimization of the automated protocol script using a CPT gradient gave 61% cell recovery. No significant differences in quality, quantity, and WBC subpopulation distribution were seen between the two freezing methods, and Mr. Frosty was selected. The manual and automated protocols were reproducible in terms of quantity, recovery, viability, WBC subpopulation distribution, gene expression, and LCL transformation. Most (75%–100%) of the 13 robustness parameters were accepted for both methods with an 8 h pre-centrifugation delay versus 38%–85% after 24 h. Differences identified between the automated and manual methods were not considered consequential.

Conclusions:

We validated the first fully automated method for isolating viable PBMCs, including RNA analysis and generation of LCLs. We recommend processing within 8 h of blood collection.

Introduction

We present here the fourth in a series of articles on the optimization and validation of biospecimen processing methods for downstream applications.^1–3 For biobanks, this is an important aspect of quality assurance, particularly in the context of automated handling, and is a normative requirement for biobank accreditation.^4,5 Optimization and validation are based on fitness-for-purpose, collection and processing efficiency, reproducibility, and robustness.

Peripheral blood mononuclear cells (PBMCs), blood cells with a rounded nucleus, include a diverse population of cells participating in the body's immune defense, notably lymphocytes (T cells, B cells, and natural killer cells), monocytes, and dendritic cells. PBMCs can be used for a broad range of downstream applications. Sorted PBMC subpopulations are increasingly being used for identification of clinically relevant biomarkers. Examples include lymphocyte subset-specific gene expression signatures in cancer⁶ or autoimmune diseases,⁷ lymphocyte subset-specific miRNA signatures in multiple sclerosis,⁸ or T cell subset-specific flow cytometric signatures in Parkinson's disease.⁹ Viable PBMCs are sorted for specific subpopulations, via immunomagnetic sorting to obtain purified monocyte and lymphocyte populations,¹⁰ and are used for functional studies,¹¹ immunophenotyping,¹² establishment of lymphoblastoid cell lines (LCL),¹³ and purification of CD34+ cells.¹⁴ Functional assays for monitoring immune responses are often used in vaccine development.

Different white blood cell (WBC) types can be identified on the basis of their expression of CD antigens. CD45 is the most widely expressed antigen and is found on all WBCs. CD3, CD14, and CD15 are characteristic of T lymphocytes, monocytes, and granulocytes, respectively. WBC types have specific gene expression signatures¹⁵ and methylation profiles,¹⁶ and the use of specific subpopulations may increase resolution in the quest for identification of clinically-relevant biomarkers. Transformation of peripheral B lymphocytes by Epstein Barr virus (EBV) is the method of choice for generating LCLs.¹⁷ LCLs serve a variety of purposes including antibody generation,¹⁸ antigen presentation for immunologic assays and cancer vaccines,^19,20 and also express genes from a wide range of metabolic pathways specific to individuals from whom the cells are derived, making them an important source of material for genetic and functional research.²¹ In the context of downstream applications, high cell yields, subpopulation composition, and cell viability are the most relevant quality attributes.

Best Practices²² and Standard Operating Procedures^22–24 for processing biobank samples have been published, but formal method validation is lacking. It has been shown that anticoagulant type, time from venipuncture to PBMC isolation and cryopreservation, and isolation method may impact the performance of T cells in downstream immunological assays.^25,26 The gold standard of PBMC isolation is density gradient centrifugation, allowing separation of PBMCs from granulocytes, platelets, and red blood cells (RBCs). A number of separation media are available, of which Ficoll, a neutral, high molecular weight, hydrophilic polysaccharide, is the most widely used. Upon centrifugation, PBMCs remain at the less dense, upper interface. Separation can be done manually or using prefilled tubes, such as vacuum-driven BD Vacutainer^® Cell Preparation Tubes (CPT)™ or Greiner Bio-One Leucosep^® tubes, in which blood is separated from the density medium with a porous membrane.

The current study was designed to identify the optimal protocol for extracting PBMCs from peripheral blood in collection tubes supplemented with anticoagulant. Processing was optimized in terms of initial whole blood centrifugation conditions, choice of gradient, and PBMC freezing method. In light of the labor-intensive nature of gradient separation (a major source of inter-operator variability) alongside the advantages of automating biobanking sample processing (allowing high throughput and reducing processing time), we established an automated workflow, and compared automated and manual methods. Optimal protocols were validated with respect to fitness-for-purpose for downstream applications, based on reproducibility and robustness for pre-centrifugation delay. Parameters evaluated included cell counts, viability, and recovery, WBC subpopulation distribution, mRNA and miRNA relative quantity, and the establishment of LCLs.

Materials and Methods

Study design

The study was performed in two steps. In the initial optimization step, four parameters were evaluated; 1) centrifugation speed of whole blood (1000 g, 2000 g, 4000 g) prior to gradient separation, evaluated in terms of cell viability and WBC subpopulation distribution; 2) gradient type (Ficoll, prefilled Leucosep tubes, prefilled CPT tubes), evaluated in terms of PBMC yield (total counts), viability, and WBC subpopulation distribution; 3) freezing method (Controlled Rate Freezer [CRF], Mr Frosty [MRF]) evaluated in terms of cell yield (total counts), viability, WBC subpopulation distribution, and establishment of LCLs; and 4) automated method approach to streamline the procedure and improve operational efficiency and reproducibility, evaluated in terms of recovery. Evaluation of centrifugation, gradient, and automated versus manual optimization parameters was performed in freshly extracted PBMCs (without a freeze-thaw cycle).

Optimized automated and manual protocols were then validated in parallel. Reproducibility and robustness (4 hours vs. 8 and 24 hours pre-centrifugation delay) were evaluated according to yield (total counts and CBC), viability, recovery, WBC subpopulation distribution, gene expression, and establishment of LCLs. The reproducibility acceptance criterion was a CV <20% between samples from a given donor for all assays. The robustness acceptance criteria were assay outcomes at 8 and 24 h within 15% of the 4-h outcome, or that all samples used to establish lymphoblastoid cell lines had the same transformability.

An ongoing stability study with annual re-testing to evaluate the effect of long-term storage on fitness-for-purpose is planned. Only baseline results are included in this report (comparison of automated vs. manual processing).

Validation assays (reproducibility and robustness) were performed after PBMC isolation and cryopreservation, with the exception of CBC counts that were performed in whole blood and in freshly isolated PBMCs, prior to freezing. Two vials were frozen; one was used for cell counts, viability, and WBC subpopulation analyses, and the other for EBV transformation and RNA extraction.

Finally, variation between the automated and manual extraction methods was determined according to fitness-for-purpose in terms of PBMC yield (evaluated by CBC), recovery, viability, WBC subpopulation distribution, gene expression analysis, and establishment of LCLs.

Peripheral blood samples were collected from healthy volunteers who provided written informed consent (CNER approval ##201107/02). An overview of the workflow is shown in Figure 1.

FIG. 1.

Overview of peripheral blood mononuclear cell (PBMC) optimization and validation.

PBMC isolation protocol

Blood collection

For centrifugation optimization, three 10-mL blood samples were collected from one donor in K2E Vacutainers® containing K2EDTA (BD #367525). For gradient optimization, three donors each provided two blood samples; 10 mL in a K2E Vacutainer containing EDTA (BD #367525) for Ficoll and Leucosep, and 4 mL in a 4-mL CPT Vacutainer containing sodium citrate (BD #362760). For freezing optimization, PBMCs isolated from the gradient optimization were used. For automation optimization, one donor provided four blood samples in 8-mL CPT Vacutainer tubes (BD #362761).

For reproducibility and stability assessments, one donor provided three 10-mL blood samples in K2E Vacutainers (diluted 1:1 with PBS), and three 8-mL samples in CPT Vacutainers for each experiment. For robustness, three donors each provided three 10-mL blood samples in K2E Vacutainers (diluted 1:1 with PBS) and three 8-mL samples in CPT Vacutainers, collected within 30 min of each other. Variation between the automated and manual extraction methods was evaluated using results from evaluations for the reproducibility, robustness and stability.

All samples (except those collected for pre-centrifugation delay) were processed within 4 h of collection and maintained at room temperature (18°–24°C).

Blood processing and gradient separation

For centrifugation optimization, tubes were centrifuged at 1000 g, 2000 g, or 4000 g for 20 min, medium brake. Plasma was removed and the pellet was resuspended in an equal volume of PBS, loaded on a Ficoll-Paque PLUS gradient (density 1.077±0.001; General Electric; #17-1440-02) in a 15-mL tube (2:1), then centrifuged for 30 min at 400 g, soft brake. The interphase was collected and transferred to a 15-mL tube containing 10 mL PBS then centrifuged at 200 g for 10 min, soft break. The pellet was suspended in 1 mL PBS and divided between two tubes.

For gradient optimization, three gradient systems were compared: 1) Ficoll; 8 mL diluted blood (4 mL whole blood, collected in K2E Vacutainer, plus 4 mL PBS) was loaded on a Ficoll-Paque PLUS gradient, centrifuged as described above, and the interphase was transferred to a 15-mL tube. 2) Leucosep; 8 mL diluted blood (4 mL whole blood, collected in K2E Vaculatainer, plus 4 mL PBS) was added to a Leucosep tube (prefilled with Leucosep™ separation medium; Greiner Bio-one; #163288) and centrifuged for 10 min at 1000 g, medium brake. The top layer was removed to 5 mm above the interphase, then all liquid down to the porous barrier was transferred to a 15-mL tube. 3) CPT; 4-mL blood sample collected directly in a 4-mL tube was inverted 10 times then centrifuged for 20 min at 1800 g, medium brake. Approximately half the plasma was aspirated and the cell layer was collected and transferred to a 15-mL tube. 15-mL tubes from the three methods were topped up to 10 mL with PBS and centrifuged for 10 min, 200 g, soft brake. Supernatant was removed and pellets were resuspended in 5 mL PBS. For reproducibility, robustness and stability, samples were manually processed using the Ficoll gradient.

Cell pellets were slowly suspended in pre-chilled Cryostor CS10 cryopreservation medium (BioLife solutions; #210102) according to the manufacturer's instructions at a concentration of 5–10 x 10⁶ cells/mL and frozen in 1-mL aliquots.

Automated PBMC extraction protocol

An automated protocol for PBMC and plasma extraction was implemented with a Tecan Freedom EVO automated liquid handler with EVOware standard software (v2.5 Service Pack 3 patch 1) using a Ficoll gradient in CPT tubes and gradient centrifugation for 20 min at 1800 g, high brake. The in-house protocol was optimized to improve recovery and reproducibility in terms of; 1) detection of the liquid height in the tube after the first centrifugation, 2) calculation of the separation gel height based on the whole volume detection, 3) aspiration of a maximum volume of the PBMC layer without touching the gel, 4) resuspension of the cell pellet after centrifugation in 11 mL PBS, and 5) aspiration of the maximal amount of resuspended pellet to a cryotube. PBMC recovery was evaluated as the total number of PBMCs in various fractions obtained during processing, compared to the total number in the volume loaded. Acceptance was based on a 50% PBMC recovery rate in the final cryotube.

The optimized script was used for reproducibility, robustness, and baseline stability experiments,. Pellets were slowly resuspended in pre-chilled 1.5 mL Cryostor CS10 in FluidX cryotubes (#65-7502) according to the manufacturer's instructions then frozen.

Freeze-thawing

For optimization, PBMCs pellets were resuspended in 6 mL freezing medium (RPMI1640 [Invitrogen, #72400-021] supplemented with 20% FBS HI [Invitrogen #26140] and 10% DMSO [Sigma; #D2438]), and aliquoted into six cryovials [FluidX; #65-7502] 1 mL/vial. Two freezing methods were evaluated using three vials per method. 1) Controlled Rate Freezer (CRF); vials were placed in an IceCube 14S Controlled Rate Freezer (Sy-Lab) using IceCube Software v1.21a, running a program of 1°C/min cooling until −80°C, then quickly transferred to liquid nitrogen; 2) Mr. Frosty (MRF); vials were placed in a Mr. Frosty Freezing Container (Nalgene; #5100-0001) filled with isopropanol, cooled slowly overnight to −80°C, then transferred to liquid nitrogen. Vials were thawed rapidly (2 min at 37°C) 6 months later.

For validation of reproducibility, robustness and baseline stability, the Mr. Frosty method was used. PBMCs were frozen in cryovials (two aliquots per sample for reproducibility and robustness, 11 aliquots for stability) with 1 mL Cryostor CS10 per vial, placed in a Mr. Frosty Freezing Container filled with isopropanol precooled at 4°C, left for 4 h at −80°C, then transferred to the liquid nitrogen vapor phase.

Vials were thawed rapidly (2 min at 37°C), then washed with 9 mL RPMI1640 supplemented with 10% FBS. Cells were centrifuged at 300 g for 5 min at room temperature, supernatant was removed and the pellet was resuspended in 1 mL RPMI-10 for evaluation assays.

Evaluation assays

Complete blood counts (CBCs)

A 100-μL sample of whole blood was counted on an ABX Micros CRP 200 hematology counter (Horiba), according to the manufacturer's instructions. Total PBMC was calculated by multiplying the sum of lymphocytes and monocytes per mL times the blood volume used.

PBMC cell count by CASY

PBMCs were quantified on a CASY Cell Counter and Analyzer (Roche Applied Science) using an in-house PBMC counting protocol (unpublished validation data). Counted samples were diluted to be within the reliable counting range. For reproducibility, recovery was evaluated by 1) comparing the total PBMC counts (CASY) after isolation relative to total PBMCs (ABX lymphocyte plus monocyte counts) in whole blood, and 2) comparing total PBMC counts (CASY) before and after freezing. Total PBMC yield was calculated by counted events/mL, times the dilution, times the total volume.

Cell viability

Cell viability was assessed by flow cytometry analysis using the FITC Annexin V Apoptosis Detection Kit 1 (BD Pharmingen; #556547) according to the manufacturer's instructions on a BD FACSVerse with FACSuite v 1.0.4.2650 software. In this article, single Annexin V-stained cells are reported as “early apoptotic”, and double Annexin V / propidium iodide-stained cells are reported as “late apoptotic” cells.

WBC subpopulation distribution

WBC distribution was assessed by flow cytometry using multiplex marking of PBMCs with a panel of fluorescently labeled monoclonal antibodies (CD45∼FITC, BD #555482; CD3∼PerCP, Miltenyi #130-094-965; CD14∼PE, BD #345785; CD15∼PECy7, BD #560827). Stained samples were analyzed on a BD FACSVerse using FACSuite v 1.0.4.2650 software.

Establishment of lymphoblastoid cell lines

Approximately 2–5×10⁶ PBMCs were exposed to EBV (supernatant from B95-8 cell line, ATCC) for 60 to 90 min at 37°C. Cells were then washed and resuspended in RPMI-20 supplemented with 1 μg/mL cyclosporine A (Sigma #C1832). Cells (∼2 x 10⁶/mL) were plated in 96-well plates and cultured at 37°C, 5% CO_2, in a humidified environment. Transformation was evaluated after 3 weeks in terms of clump formation score.

RNA extraction and spectrophotometry

Total RNA, including miRNA, was extracted from 2–5×10⁶ PBMCs using the miRNeasy mini kit (QIAgen #217004) on a QIAcube then quantified by spectrophotometry on a Synergy MX (Biotek). To be in the range for downstream PCR applications, eluted RNA samples were concentrated using a CentriVap (Labconco), resuspended in 15 μL PCR grade water and quantified again.

mRNA and miRNA qRT-PCR

mRNA (hDUSP1, hJUN) and miRNA (RNU24, miR16) were quantified using commercial TaqMan assays (Life Technologies) according to the manufacturer's instructions using commercial probes(Life Technologies). qPCR reactions were run on the ABI 7500 Fast Real Time PCR System (Life Technologies) and results were scored in Cts.

Statistical analyses

Mean, standard deviation (SD) and CV% were calculated using Microsoft Excel 2010. Optimization outcomes and variability between automated and manual methods were compared using paired two-tailed t-tests. Significance was calculated using Sigma Plot v.12.0 (Systat Software) with a 5% significance threshold using Tukey and Dunn tests.

Results

Optimization of PBMC isolation protocols

Centrifugation conditions

Initially, processing of whole blood was optimized in terms of centrifugation speed to determine the feasibility of simultaneously isolating plasma and PBMCs from a single sample, based on our previously validated method for plasma isolation.¹ The majority of PBMCs isolated from whole blood were viable, and similar proportions were isolated with all three speeds following Ficoll gradient separation; 91.7% at 1000 g, 88.1% at 2000 g, and 94.0% at 4000 g (CV 3.3%). Late apoptotic cells accounted for ≤10% of isolated PBMCs, and early and necrotic PBMCs for <2%. The proportions of CD45+, CD3+, and CD14+ cells were similar with each of the three speeds (CV 1.1%, 5.2% and 6.6%, respectively; Supplementary Table S1; supplementary material is available in the online article at www.liebertpub.com/bio). A high 32.2% CV seen for CD15+ cells was probably due to the low proportions of these cells detected (<3% of cells).

Gradient separation

Three gradient separation methods were compared in terms of cell yield, viability and WBC subpopulation distribution (Table 1).Cell counts varied between the three methods, notably for two of the three donors. The Ficoll method consistently gave the highest cell yields, followed by Leucosep which gave from 5% to 17% fewer cells, while the CPT method gave from 13% to 50% fewer cells than the Ficoll method. In terms of cell viability, CPT gave 9% to 13% fewer viable cells than either the Ficoll or Leucosep method for two donors and was approximately equivalent for the third, with CV% ranging from 1.2% to 7.7%. Very little difference was seen between the three methods in terms of WBC subpopulation distribution, with <5.5% CV for leukocytes (CD45+), lymphocytes (CD3+) and monocytes (CD14+). PBMCs isolated with Leucosep and CPT had lower granulocyte contamination (CD15+) compared to Ficoll. The CV% was much higher with CD15+, likely due to the very small percent positive cells (<4%). Ficoll was selected as the gradient for manual PBMC isolation for subsequent reproducibility and robustness evaluations.

Table 1.

Comparison of Manual Gradient Separation Methods of Peripheral Blood Mononuclear Cells (PBMCs) in Terms of Yield, Viability, and White Blood Cell (WBC) Composition

		Ficoll (EDTA)	Leucosep (EDTA)	CPT (citrate)	Mean^a (SD)	CV%
PBMC counts (CASY)
Donor 1	Total PBMCs (×10⁶)^b	7.05	6.71	6.12	6.6 (0.47)	7.1%
Donor 2	Total PBMCs (×10⁶)^b	9.60	8.94	6.37	8.3 (1.7)	20.6%
Donor 3	Total PBMCs (×10⁶)^b	10.15	8.40	5.54	8.0 (2.3)	29.0%
Viability (FACSVerse)
Donor 1	Viable cells (%)	92.9	92.1	90.7	91.9 (1.1)	1.2%
Donor 2	Viable cells (%)	92.4	92.7	80.8	88.6 (6.8)	7.7%
Donor 3	Viable cells (%)	94.2	92.4	85.8	90.8 (4.4)	4.9%
WBC composition
Donor 1	CD45+ (%)	93.8	99.2	98.8	97.3 (3.0)	3.1%
	CD3+ (%)	63.7	63.8	62.8	63.4 (0.6)	0.9%
	CD14+ (%)	15.7	16.9	17.4	16.7 (0.9)	5.1%
	CD15+ (%)	1.0	0.5	0.6	0.7 (0.3)	41.2%
Donor 2	CD45+ (%)	97.1	98.4	99.4	98.3 (1.2)	1.2%
	CD3+ (%)	62.3	65.3	63.0	63.5 (1.6)	2.5%
	CD14+ (%)	15.9	16.3	16.5	16.2 (0.3)	1.9%
	CD15+ (%)	1.3	0.3	0.4	0.7 (0.5)	77.8%
Donor 3	CD45+ (%)	91.9	91.7	88.9	90.8 (1.6)	1.8%
	CD3+ (%)	59.6	64.8	66.0	63.4 (3.4)	5.4%
	CD14+ (%)	12.1	11.9	11.9	11.9 (0.1)	1.0%
	CD15+ (%)	3.9	0.4	0.5	1.6 (2.0)	123.4%

Mean of the three methods per donor; ^bTotal PBMCs=Counts for 1 mL × total volume (5 mL).

Automation

CPT, which is compatible for use with the Tecan automated liquid handling system, was used for gradient separation. Recovery of total PBMCs was evaluated in various fractions obtained with the automated method and compared with whole blood counts for two successive optimization protocols (Table 2). After the first (non-optimized) script, a 15% yield was obtained in the final aliquots. The greatest sources of cell loss were the volume left above the gel after PBMC aspiration (to avoid touching interphase) and the volume left in the mixing tube after aspiration. A first script optimization was performed taking these aspects into account, which resulted in a 42% yield; however, this was still below the required 50% recovery threshold. Further modifications were implemented (second optimization) to improve the yield, moving the liquid aspiration to a lower level and using a larger volume pre-aspiration mix. This final revised script gave a 61% yield and was used for validation.

Table 2.

Optimization of Automated Peripheral Blood Mononuclear Cell (PBMC) Isolation According to Two Scripts (Tecan Liquid Handler) in Terms of PBMC Yield

	Initial script^a			Optimization 1^b			Optimization 2^c
Fraction	Volume (mL)	Total PBMCs (10⁶)	% total PBMC	Volume (mL)	Total PBMCs (10⁶)	% total PBMC	Volume (mL)	Total PBMCs (10⁶)	% total PBMC
Whole blood^d	9	18	100%	9	18	100%	9	11.2	100%
Volume above gel after PBMC aspiration	0.7	4.62	26%	0.35	2.17	12%	nd	nd	nd
Washing gel with 1 ml PBS	1	1.4	8%	1	2.3	13%	nd	nd	nd
Volume left in mixing tube after aspiration	0.3	4.23	24%	0	0	0	0	0	0%
Final cryotube aliquot	1.5	2.7	15%	1.6	7.52	42%	1.6	6.75	60.9%

Initial (non-optimized) protocol script; ^bFirst revised protocol script; ^cSecond revised protocol script; ^dBlood diluted with 2.5 mL citrate; nd, not determined.

Freeze-thawing

The effect of two freezing methods, the Controlled Rate Freezer and Mr. Frosty, on cell yield, viability, WBC subpopulation distribution, and biological activity was compared for three donors. No significant differences between the two methods were seen for total PBMC or WBC subpopulation distribution for any of the three donors (Table 3). Some fluctuations were apparent in subpopulation distribution, with Mr. Frosty favoring lymphocytes and Controlled Rate Freezer favoring monocytes (data not shown). A significant difference between the two methods in terms of the number of viable cells was identified in one of the three donors, however overall, post-thaw viability was high (approximately 80%). WBC subpopulation distribution was not significantly different between the two freezing methods for any of the donors. Finally, PBMCs from all three donors were 100% compatible with EBV transformation for both freezing methods. The Mr. Frosty method was applied for subsequent reproducibility and robustness evaluations.

Table 3.

Optimization of Peripheral Blood Mononuclear Cell (PBMC) Freezing Methods in Terms of Yield, Viability, and White Blood Cell (WBC) Subpopulation Distribution

	Donor 1^a			Donor 2^a			Donor 3^a
	Mean (SD)			Mean (SD)			Mean (SD)
	CRF	MRF	P value^b	CRF	MRF	P value^b	CRF	MRF	P value
PBMC counts (CASY)
Total PBMCs (×10⁶)^c	1.86 (0.26)	2.04 (0.10)	0.238	2.58 (0.15)	2.52 (0.05)	0.542	2.23 (0.05)	2.18 (0.16)	0.614
Viability (FACSVerse)
Healthy cells (%)	78.57 (2.45)	83.16 (0.72)	0.128	83.18 (0.42)	86.19 (1.02)	0.036	77.88 (4.73)	82.51 (1.33)	0.242
WBC Composition
CD45+ (%)	99.03 (0.12)	99.10 (0.10)	0.635	99.63 (0.06)	99.53 (0.12)	0.423	99.37 (0.25)	99.20 (0.10)	0.199
CD3+ (%)	50.17 (2.48)	52.67 (0.32)	0.216	51.923 (0.92)	54.33 (0.40)	0.086	49.53 (4.17)	51.87 (1.36)	0.540
CD14+ (%)	24.87 (0.84)	22.60 (0.46)	0.093	23.73 (1.29)	22.40 (0.66)	0.286	19.83 (1.10)	19.93 0.64	0.930
CD15+ (%)	0.70 (0.35)	0.47 (0.06)	0.336	0.60 (0.10)	0.77 (0.21)	0.370	2.20 (0.80)	2.10 (0.36)	0.845

CRF, Controlled Rate Freezer; MRF, Mr Frosty.

Bold, p<0.05.

For each donor, cells were pooled from the three manual gradient methods; ^bTwo-tailed paired t-test; ^cTotal PBMCs=Counts for 1 mL × total volume (1 mL); single reading.

Reproducibility of manual and automated PBMC isolation protocols

The manual Ficoll (EDTA) and the automated CPT (citrate) protocols were validated for reproducibility in terms of cell quantity and quality, functionality, and gene expression, using an acceptance criterion of a CV <20% between three samples from the same donor (Table 4). With the exception of CD15+, all parameters assayed in PBMCs extracted using the manual Ficoll method were within the 20% acceptance criterion, with most parameters having a CV of approximately 5% or less. CD15+ is a granulocyte marker which should not be present in the PBMC interphase. Increased CD15+ cells indicate granulocyte contamination in the Ficoll layer. CV% variability was greater for PBMCs isolated using the Tecan automated liquid handler, with several parameters having a CV >15% including three parameters with a CV >20% (RBC and platelet counts, and PBMC pre-freeze/thaw recovery).

Table 4.

Reproducibility of Peripheral Blood Mononuclear Cells (PBMCs) According to Acceptance Criteria for Cell Yield, Viability, White Blood Cell (WBC) Subpopulation Composition, and Ribonucleic Acid (RNA) Targeted Gene Expression

		Automated (CPT)		Manual (Ficoll)
		Mean (SD)^a	Acceptance criteria CV (%)	Mean (SD)^a	Acceptance criteria CV (%)
Whole Blood
CBC counts	WBC (10⁶/mL)	5.2 (0.36)	6.9	6.3 (0.1)	1.6
	Lymphocytes (10⁶/ml)	1.2 (0.17)	14.4	1.43 (0.06)	4.0
	Lymphocytes (%)	23.5 (2.27)	9.6	24.0 (0.62)	2.6
	Monocytes (10⁶/mL)	0.2 (0)	0	0.3 (0)	0
	Monocytes (%)	4.9 (0.36)	7.4	5.4 (0.23)	4.3
	Granulocytes (10⁶/ml)	3.8 (0.2)	5.3	4.6 (0.06)	1.3
	Granulocytes (%)	71.6 (2.58)	3.6	70.6 (0.8)	1.1
	RBC (10⁹/mL)	5.1 (1.05)	20.5	5.3 (0.12)	2.3
	Platelets (10⁶/mL)	157.3 (53.05)	33.7	192.7 (9.87)	5.1
Freshly isolated PBMCs
Cell counts (CASY)^b	Total cells (10⁶/mL)	4.71 (0.94)	19.9	1.05 (0.08)	7.2
	PBMC Recovery^b (%)	60.9 (15.41)	25.3	72.1 (6.01)	8.3
Frozen isolated PBMCs
Cell counts (CASY)^c	Total cells (10⁶/mL)	3.30 (0.59)	18.0	3.90 (0.23)	5.8
	PBMC Recovery^c (%)	83.9 (3.4)	4.1	64.8 (6.3)	9.7
Cell viability, FACSVerse	Necrosis (%)	1.09 (0.22)	19.7	0.86 (0.13)	15.0
	Late apoptosis (%)	13.17 (1.50)	11.4	14.22 (0.35)	2.5
	Healthy cells (%)	75.03 (1.15)	1.5	78.71 (0.55)	0.7
	Early apoptosis (%)	10.71 (0.29)	2.7	6.21 (0.28)	4.5
WBC distribution	CD45+ (%)	98.6 (0.35)	0.4	99.2 (0.09)	0.1
	CD3+ (%)	63.8 (1.45)	2.3	58.8 (0.58)	1.0
	CD14+ (%)	14.7 (1.46)	10.0	17.2 (0.47)	2.7
	CD15+ (%)	1.6 (0.26)	15.6	2.40 (1.36)	57.4
RT-qPCR	hDUSP1 (Ct)	20.02 (1.56)	7.8	18.48 (0.88)	4.8
	hJUN (Ct)	26.34 (1.30)	5.0	24.14 (0.69)	2.9
	RNU24 (Ct)	23.63 (0.23)	1.0	23.54 (0.63)	2.7
	miR16 (Ct)	20.96 (0.25)	1.2	20.20 (1.09)	5.4
LCL transformation	100%	+	+	+	+

Bold indicates assays for which CV% was >20%.

Mean of three samples from one donor; ^bTotal PBMC counts (CASY) after isolation relative to total PBMCs (ABX lymphocyte plus monocyte counts) in whole blood; ^cTotal PBMC counts (CASY) after freezing relative to before.

For each processing method (manual and automatic), both mRNA (hDUSP1 and hJUN) and miRNA (RNU24 and miR16) C+ values showed good reproducibility with CVs <20% (CV ranged from 0.97% to 7.77%). Both methods were reproducible for the functional assay of LCL transformation, with 100% transformation in all cases. Overall, both methods were reproducible according to the acceptance criterion of CV <20%.

Robustness of manual and automated PBMC collection protocols

Cell quality, quantity, and functional parameters were evaluated for robustness by comparing an 8 h and a 24 h pre-centrifugation delay to a 4 h delay. Most parameters were accepted (i.e., were within 15% of the values at 4 h) for all donors with both methods for the 8h pre-centrifugation delay (Table 5). However when the pre-centrifugation delay was increased to 24 h, acceptance varied between donors and methods, ranging from 38% to 85%.

Table 5.

Robustness of Peripheral Blood Mononuclear Cell (PBMC) Protocol with 8 H and 24 H Pre-centrifugation Delay, According to Cell Quantity, Quality, and Biological Function, Post Freeze-Thaw

		Donor 1		Donor 2		Donor 3
		Automated (CPT)	Manual (Ficoll)	Automated (CPT)	Manual (Ficoll)	Automated (CPT)	Manual (Ficoll)
8 hour delay
CBC counts	Total cells (/ml)	acc	acc	rej	rej	acc	acc
	WBC (10⁶/ml)	rej	acc	acc	rej	acc	acc
	Lymphocytes (%)	acc	acc	acc	acc	acc	acc
	Monocytes (%)	acc	acc	acc	rej	acc	acc
Cell viability (FACSVerse)	Viable cells (%)	acc	acc	acc	acc	acc	acc
WBC distribution	CD3+ (%)	acc	acc	acc	acc	acc	acc
	CD14+ (%)	rej	acc	rej	acc	acc	acc
	CD15+ (%)	acc	rej	acc	acc	rej	acc
RT-qPCR	hDUSP1 (Ct)	acc	acc	acc	acc	acc	acc
	hJUN (Ct)	acc	acc	acc	acc	acc	acc
	RNU24 (Ct)	acc	acc	acc	acc	acc	acc
	miR16 (Ct)	acc	acc	acc	acc	acc	acc
LCL transformation		acc	acc	acc	acc	acc	acc
8 hour % accepted parameters		85	92	85	77	92	100
24 hour delay
CBC counts ABX)	Total cells (/ml)	rej	acc	rej	rej	acc	acc
	WBC (10⁶/ml)	rej	rej	rej	rej	acc	acc
	Lymphocytes (%)	acc	acc	rej	acc	acc	acc
	Monocytes (%)	rej	acc	rej	rej	rej	rej
Cell viability (FACSVerse)	Viable cells (%)	rej	rej	rej	rej	acc	rej
WBC distribution	CD3+ (%)	acc	acc	rej	rej	acc	acc
	CD14+ (%)	rej	rej	rej	rej	rej	rej
	CD15+ (%)	rej	rej	rej	rej	acc	rej
RT-qPCR	hDUSP1 (Ct)	acc	acc	acc	acc	acc	acc
	hJUN (Ct)	acc	acc	acc	acc	acc	acc
	RNU24 (Ct)	acc	acc	acc	acc	acc	acc
	miR16 (Ct)	acc	acc	acc	acc	acc	acc
LCL transformation		acc	acc	acc	acc	acc	acc
24 hour %accepted parameters		54	69	38	46	85	69

Acc, <15% of 4 h value; Rej, ≥15% variation from 4 h value.

Variation between automated and manual extraction methods

Variation between the CPT automated and the EDTA/Ficoll manual methods was determined for each of the nine samples collected per donor for reproducibility, 4 h robustness and baseline stability (three replicates each), and evaluated according to the two isolation methods. Initially, a comparison of CBC counts in whole blood collected in citric acid CPT Vacutainers and EDTA K2E Vacutainers was performed. Blood samples collected in the automated CPT tubes had approximately 17% fewer WBCs and platelets compared to Ficoll tubes, while lymphocyte, monocyte, granulocyte, and RBC counts were approximately the same with both collection tubes (Table 6).

Table 6.

Comparison of Complete Blood Cell (CBC) Counts in Isolated Peripheral Blood Mononuclear Cell (PBMC) with Automated Extraction (Citric Acid) Versus Manual Extraction (EDTA)

		Ratio of Automated/Manual (%)
		Mean (SD)^a
CBC counts	White blood cell (10⁶/mL)	82.6 (8.5)
	Lymphocytes (%)	99.1 (11.1)
	Monocytes (%)	96.1 (13.4)
	Granulocytes (%)	101.2 (6.4)
	Red blood cell (10⁹/mL)	96.3 (17.1)
	Platelets (10⁶/mL)	83.2 (23.2)

Mean (SD) percent ratio comparing automated outcome versus manual outcome for each parameter from nine samples (reproducibility, robustness and stability-related samples) for each isolation method.

Although PBMC recovery in freshly isolated samples compared to whole blood was higher with the manual method compared to the automated method (69.7% vs. 60.9%; Table 7), this finding was not statistically significant and was balanced out by the freeze/thaw cycle, with the automated method giving higher recovery after a passage in liquid nitrogen (87.4% vs. 65.4%). Significantly higher viability was found with the manual extraction after one freeze-thaw cycle (79.9% versus 74.0%; p=0.01). All data taken together, the final post-thaw yield in terms of recovery of viable PBMCs is 36.4% for the manual method and 39.4% for the automated method.

Table 7.

Variation Between Automated and Manual Peripheral Blood Mononuclear Cell (PBMC) Isolation Methods, According to Cell Quantity, Immunophenotype, Gene Expression, and Lymphoblastoid Cell Line (LCL)-Compatibility Parameters

		Automated (Citrate)		Manual (EDTA/Ficoll)
		Mean (SD)^a	CV (%)	Mean (SD)^a	CV (%)	P value
Freshly isolated PBMCs
Cell counts (CASY)	PBMC Recovery^b (%)	60.9 (13.5)	22.3	69.7 (5.7)	8.2	0.21
Frozen isolated PBMCs
Cell counts (CASY)	PBMC Recovery^c (%)	87.4 (7.9)	9.0	65.4 (14.8)	22.6	<0.01
Cell viability (FACSVerse)	Healthy cells (%)	74.0 (3.4)	4.6	79.9 (2.4)	3.0	0.01
White blood cell distribution	CD3+ (%)	60.8 (5.0)	8.2	57.9 (6.0)	10.4	0.03
	CD14+ (%)	15.3 (1.4)	9.2	17.7 (2.1)	12.1	0.02
	CD15+ (%)	1.4 (0.6)	43.8	1.3 (1.3)	94.9	0.82
RT-qPCR	hDUSP1 (Ct)	19.9 (1.4)	6.7	18.8 (1.0)	5.1	0.055
	hJUN (Ct)	25.8 (1.5)	5.6	24.1 (0.8)	3.1	0.015
	RNU24 (Ct)	20.8 (1.1)	5.2	20.6 (0.9)	4.3	0.68
	miR16 (Ct)	23.5 (0.4)	1.7	23.6 (0.5)	1.2	0.65
LCL transformation	100%	+		+

Bold, p<0.05.

Mean of three samples from three donors, stability samples were pooled; ^bAfter PBMC isolation and pre-freeze/thaw; total PBMC counts (CASY) after isolation relative to total PBMCs (ABX lymphocyte plus monocyte counts) in whole blood; ^cPost-freeze/thaw; total PBMC counts (CASY) after freezing relative to before.

Significantly different proportions of both CD3+ and CD14+ populations were present in PBMCs isolated by the automated method compared to the manual method (p=0.03 and 0.02, respectively); however, the CV% of each method were within the reproducibility acceptance criterion. From identical RNA starting quantities, a statistically significant difference was seen in hJUN mRNA quantification, with the automated method yielding significantly less hJUN than the manual method (p=0.015) but not in hDUSP, while no statistically significant differences were found for miRNA target quantification. This variation may be due to hJUN expression being sensitive to the citrate anticoagulant in the CPT samples. There was no difference between the two extraction methods in terms of transformability of LCLs.

Discussion

The value of automated isolation of PBMCs over manual Ficoll separation was first reported for cell therapy purposes in 2008,²⁷ confirming significantly higher recovery of clinical grade PBMCs using a Sepax system, while also eliminating inter-user differences. Here we report for the first time automated isolation of PBMCs for research purposes using a current widely available liquid handler.

The initial optimization step demonstrated the feasibility of simultaneously obtaining plasma and PBMCs from a single whole blood sample with Ficoll manual separation in terms of viable cells and WBC subpopulation distribution. The 2000 g speed was selected as optimal on the basis of a previous report for validation of plasma isolation,¹ and is the recommended centrifugation speed for obtaining both plasma and viable PBMCs suitable for downstream research applications from a single blood sample.

Of the three manual gradient methods, Ficoll and Leucosep gave higher PBMC yields than CPT. Besides different anticoagulant and processing methods, this may be attributed to a potentially smaller starting volume of blood due to CPT tubes allowing a maximum 4-mL sample, leading to potentially smaller collected volumes, whereas the other two methods guaranteed a 4-mL starting volume. Both Ficoll and Leucosep outperformed CPT in terms of pre-freeze PBMC viability. Nonetheless, the CV across the three gradients was less than 20% for viability and all WBC subpopulations other than granulocytes. CPT and Leucosep both gave less granulocyte contamination than Ficoll. Published data comparing gradient methods are conflicting, with Schlenke et al. reporting higher PBMC yields with CPT compared to Ficoll but lower viability,²⁸ while Nilsson et al. showed the inverse in one institution and equivalence in another institution.²⁹ Ruitenberg also reported equivalence in terms of viability and recovery.³⁰ In our study, with the exception of yield, the choice of gradient did not appear to influence the outcome, and all three separation methods were considered fit-for-purpose given the high yields. With freezing PBMCs for long-term storage being the main purpose of this procedure, from a fit-for-purpose perspective, both the CPT and Ficoll methods are appropriate. Ficoll was selected for further evaluations of the manual method, as the most widely implemented method and providing the highest yields.

Although mainly used post-cryopreservation, it has been reported that viability of Ficoll-derived PBMCs from cryopreserved samples is significantly lower than in freshly isolated samples.³⁰ Nonetheless, the significance of this difference appears to be of little consequence in our experimental setting, as viability of both fresh and frozen PBMCs was 90% or more. We found that lymphocyte recovery was lower in frozen whole blood than frozen PBMCs isolated from the same source (unpublished data), although it has been reported that lymphocyte distribution (CD4+ and CD8+) is not significantly different between them.³¹ We thus recommend that if recovered quantity is important, freezing of extracted PBMCs is preferred over whole blood. Furthermore, post-thaw viability was found to be higher in PBMCs isolated with the automated method compared to the manual method. PBMCs isolated by the automated method have been exposed to Cryostor CS10 solution, potentially compromising the immediate post-isolation quality, but improving recovery after a freeze-thaw cycle.

Although freezing of isolated PBMCs is an essential aspect of the use of PBMCs in biobanking, few reports evaluating freezing methods have been published. Buhl et al. compared CRF freezing with standard isopropyl alcohol freezing for downstream dendritic cell applications.³² They showed that while similar dendritic cell viability was seen with the two methods, induction of antigen-specific IFN-γ release from autologous effector T cells was significantly higher with CRF. In our study, the two freezing methods, Mr. Frosty and Controlled Rate Freezer, gave largely similar cell recovery, viability, cell subpopulation distribution, and susceptibility to EBV immortalization. Mr. Frosty was selected as the preferred method in light of ease of handling and administration.

Short-term stability studies support the added value of using Cryostor CS10 cryopreservation medium.³³ We have initiated a long-term (8-year) stability study comparing PBMCs isolated by the CPT automated and Ficoll manual methods. Both PBMCs and whole blood have previously been shown to be stable in terms of lymphocyte subpopulation distribution after cryopreservation at −80°C for 10 or 120 days.³¹ Schelenke showed significant increases in granulocyte contamination with longer storage time following Ficoll isolation, but this parameter was stable with CPT.²⁸ Recovery and viability of PBMCs cryopreserved for up to 12 years in biobanks has been reported.³⁴ However, long-term cryopreservation of PBMCs has been shown to have a negatively significant effect on CD4+ T cell responses to different antigens, possibly due to increased CD4+ T cell apoptosis.³⁵ Therefore, for functional assays, we recommend that case and control samples have similar “storage ages”.

Both the automated and manual PBMC extraction methods were considered reproducible, with the acceptance criterion of CV<20% fulfilled for almost all quantitative parameters. Although RBC and platelet counts were outside the CV% criterion with the automated method, neither are critical quality attributes of PBMCs and this variation is likely attributed to the difference in anticoagulant in the CPT and K2EDTA Vacutainers (citric acid and EDTA, respectively). The non-reproducibility for CD15+ distribution with the manual method may be due to contamination of the Ficoll layer by these cells that are then aspirated together with the PBMCs. We concluded that both the automated and the manual PBMC extraction methods are reproducible.

It has been shown that the length of time from venipuncture to cryopreservation is the most critical parameter affecting the performance of T cells in downstream immunological ELISpot assays,²⁶ and also affects the cytotoxic activity of natural killer cells.³⁶ It has also been shown that a 24-h delay between blood collection and processing, especially at 4°C, significantly increases granulocyte contamination, which may inhibit T cell function.³⁷ All of these studies recommend a maximum delay of 8 h at room temperature. Likewise, Mallone et al. reported that whole blood should ideally be processed as soon as possible after being drawn, within a maximum of 8 to 12 h or diluted in medium to minimize granulocyte contamination and preserve T-cell response.³⁸ We found that neither extraction method was 100% robust for all parameters with an 8-h or a 24-h pre-centrifugation delay. Nonetheless, most parameters were acceptable with an 8-h delay, which is suitable for the majority of assays. Ribosomal RNA integrity, as measured by RNA integrity number (RIN), is not significantly altered after 24 h at room temperature and is largely unaffected at earlier time points.³⁹ The two “universal” miRNA examined in our study, miR16 and miR24, were also stable with pre-centrifugation delays of up to 24 h. DUSP1 and hJUN have been reported to be unstable with short precentrifugation delays,⁴⁰ and while DUSP1 was largely stable in our study, JUN showed a slight increase over 24 h (Supplementary Table S2; supplementary material is available in the online article at www.liebertpub.com/bio). These results suggest that various PBMC mRNAs may be down- or upregulated by prolonged transport times.

PBMCs isolated with the automated method were comparable overall in terms of quantity and quality post-freeze/thaw to manually extracted PBMCs isolated with Ficoll. The variations in viability and recovery balanced each other out, with the 20% higher recovery for automated extraction versus 5% higher viability with manually processed samples. The approximately 17% lower WBC and platelet counts with CPT tubes may have been impacted by the different anticoagulants used with the two methods. Other variations, such as CD3+ and CD14+ distribution, were not considered to have a major impact in the context of downstream applications, given that the differences were within the reproducibility acceptance criterion. Minor variations in RNA gene expression levels were observed with the two methods, however this was only of statistically significance for the hJUN gene. Nonetheless, this is difficult to interpret given that mRNA from PBMCs isolated from blood treated with different anticoagulant cannot be compared.

While the final automated protocol achieved adequate recovery, further improvement will be difficult because of the risk of clogging if the tip touches the separating gel when aspirating. The automated protocol provides a safety margin in terms of the position of the gel that was defined in the script employed in this study, as a constant percentage of the total liquid volume. However, given the wide hematocrit heterogeneity in the general population (normal values from 37%–52%), the algorithm defining the gel position needs to be able to accommodate a wide range of blood samples, and as such, mean recovery will not increase above 60% with currently technology.

Automated isolation of viable PBMCs using CPT tubes is feasible, reproducible, and robust, giving good yield, recovery, and post-thaw viability. It also has comparable WBC subpopulation composition to PBMC populations manually isolated with EDTA/Ficoll, with all cell subpopulations being present. The absence of depletion of a specific subpopulation eliminated the likelihood of bias. We present our method as a recommended protocol for automated processing of whole blood to obtain PBMCs for research purposes (Table 8). Automated processing greatly improves logistics in biobanks processing large numbers of samples for downstream assays requiring viable PBMCs, such as in the context of clinical trials. Samples should be processed within 8 hours of blood collection. It is essential that methods be thoroughly reported^41,42 to ensure comparability of outcomes (collection tube type, pre-centrifugation delay, etc).

Table 8.

Method Overview for Automated Peripheral Blood Mononuclear Cell Isolation from Whole Blood

Parameter	Blood sample
Primary container type	CPT citrate tube 8 ml
Pre-centrifugation delay between collection and processing	Maximum 8 hours at room temperature
Gradient centrifugation	1800 g; 20 min; room temperature; high brake force
Automated liquid handling	Collect cells, wash, suspended in pre-chilled Cryostor CS10, and aliquot
Final container type	2 ml FluidX Cryotube
Freezing	Mr Frosty isopropanol, slow overnight cooling
Long-term storage	Liquid nitrogen

Footnotes

Acknowledgments

We thank Sarah MacKenzie, PhD for medical writing assistance, and Hans Peter Sattler, Matthias Greuter, and Jonas Loof from Tecan for excellent technical assistance.

Author Disclosure Statement

No competing financial interests exist.

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