Highly sensitive in vitro cytokine release assay incorporating high-density preculture

Abstract

Immunostimulatory effects of monoclonal antibodies (mAb) through binding to F_cγ receptors (F_cγR) on immune cells are a likely cause of cytokine release syndrome. However, it is difficult to detect the potential risk of F_cγR-dependent cytokine release associated with mAb in the current standard cytokine release assays (CRA), including the air-drying solid-phase method using human peripheral blood mononuclear cells (PBMC). To increase the sensitivity to detect F_cγR-dependent cytokine release due to mAb, a high-density preculture (HDC) method was incorporated into the air-drying solid-phase CRA. Here, PBMC were exposed to panitumumab, trastuzumab, rituximab, or alemtuzumab at 0.1, 0.3, 1, and 3 μg/well for 24 or 48 hr under both non-HDC and HDC conditions. T-cell agonists (anti-CD3 mAb, anti-CD28 super-agonist [SA] mAb) were used as reference mAb. Panitumumab, trastuzumab, rituximab, or alemtuzumab induced cytokine release under both non-HDC and HDC conditions, and cytokine release caused by alemtuzumab was more pronounced under HDC conditions. To investigate F_cγR involvement in cytokine release associated with panitumumab, trastuzumab, rituximab, and alemtuzumab, CRA of these four mAb were conducted with anti-F_cγRI, -F_cγRII, or -F_cγRIII F(ab’)₂ fragments. The results showed cytokine release caused by trastuzumab, rituximab, and alemtuzumab was significantly suppressed by anti-F_cγRIII F(ab’)₂ pretreatment, and slightly reduced by anti-F_cγRI or anti-F_cγRII pretreatment, indicating these mAb induced F_cγR (especially F_cγRIII)-dependent cytokine release from PBMC. Cytokine release caused by panitumumab was slightly suppressed by anti-F_cγRIII F(ab’)₂ pretreatment. Anti-CD3 mAb and anti-CD28 SA mAb also induced significant release of cytokines under HDC conditions compared with that under non-HDC conditions. In conclusion, CRA incorporating HDC into the air-drying solid-phase method using human PBMC could sensitively capture the F_cγR-dependent cytokine release potential of mAb.

Keywords:

• Cytokine release syndrome
• high-density preculture
• F_cγR
• cytokine release assay
• therapeutic monoclonal antibody
• peripheral blood mononuclear cell

Introduction

Cytokine release syndrome (CRS), which occurred in the Phase-I clinical trial of the anti-CD28 super-agonist (SA) monoclonal antibody (mAb) TGN1412, is one of the most serious adverse effects associated with parenteral administration of therapeutic mAb (Suntharalingam et al. 2006). CRS is typically characterized by the release of pro-inflammatory cytokines including interleukins (IL), interferon (IFN)-γ, and tumor necrosis factor (TNF)-α into the blood, resulting in fever, chills, hypotension, and multiple organ failure immediately after treatment with a therapeutic mAb (Finco et al. 2014). The cytokine release caused by therapeutic mAb is considered to be mediated by two mechanisms: the binding of the fragment antigen-binding region (F_ab) of mAb to the cell surface target on immune cells and the binding of the fragment crystallizable region (F_c) of mAb to F_cγ receptors (F_cγR) on effector cells (Wing et al. 1996). The former (i.e., binding of F_ab) is primarily associated with CRS associated with T-cell agonists such as OKT3 [an anti-CD3 mAb] and TGN1412; the latter (i.e., binding of F_c) is mainly associated with CRS that arises with alemtuzumab, trastuzumab, and rituximab and is due to antibody-dependent cellular cytotoxicity (ADCC) activity (Kirton et al. 2011; Paul and Cartron 2019).

Cytokine release assays (CRA) using human peripheral blood mononuclear cells (PBMC) have become common in vitro assay systems to predict the risk of clinically relevant CRS because in vivo and in vitro animal studies failed to predict this risk (Stebbings et al. 2007; Eastwood et al. 2010; Hansel et al. 2010; Pallardy and Hunig 2010). Several different assay platforms have been introduced as nonclinical risk assessment systems, including liquid-phase, wet-coating solid-phase, and air-drying solid-phase methods (Findlay et al. 2010; Römer et al. 2011; Eastwood et al. 2013). The solid-phase method is commonly used to detect the risk of CRS due to mAb (Finco et al. 2014). In fact, TGN1412 caused profound cytokine release from PBMC in the solid-phase method, but not in the liquid-phase method (Findlay et al. 2010, 2011). However, the cytokine release inducible by TGN1412 was reported to be detected using high-density precultured (HDC) PBMC even in the liquid-phase method because the HDC method enhanced T-cell reactivity via contact with monocytes (Bartholomaeus et al. 2014; Hussain et al. 2015).

Recently, CRA have been applied to evaluate the mAb that act on F_cγR in addition to the mAb having agonistic activity. However, unlike the case with TGN1412, alemtuzumab caused TNFα and IFNγ in the liquid-phase, but not in the solid-phase, method when compared with levels of each induced with an isotype control antibody (Vessillier et al. 2015). Further, the increase in TNFα due to alemtuzumab in the solid-phase method was comparable to that attained with trastuzumab or bevacizumab which have been associated with a moderate/low CRS incidence in humans (Chung 2008). Such results indicate the disadvantage of the solid-phase method. Thus, it is difficult to detect potential risks of CRS associated with mAb such as alemtuzumab that act on F_cγR in the solid-phase method (Findlay et al. 2010; Eastwood et al. 2013; Grimaldi et al. 2016).

To establish a solid-phase assay system for detecting the potential risk of CRS through the binding of mAb to F_cγR on effector cells, this study attempted to incorporate HDC-PBMC into the air-drying solid-phase method. In the studies here, several mAb, for example, trastuzumab, rituximab, and alemtuzumab were evaluated as each was reported to cause F_cγR-dependent cytokine release (Kirton et al. 2011; Paul and Cartron 2019). In addition, anti-CD3 mAb and anti-CD28 SA mAb with T-cell agonistic activity were evaluated. Panitumumab was used as a negative control because its risk of CRS has been reported to be low (Bugelski et al. 2009). Lastly, the current study also examined the types of F_cγR that contribute to the cytokine release caused by these mAb in this assay system.

Materials and methods

Antibodies and reagents

The following mAb were used in the present study: panitumumab (Vectibix^®, Amgen, Thousand Oaks, CA, USA; anti-epidermal growth factor receptor (EGFR) human IgG₂κ), rituximab (Rituxan^®, Genentech, South San Francisco, CA, USA; anti-CD20 chimeric murine/human IgG₁κ), trastuzumab (Herceptin^®, Genentech; anti-HER2 humanized IgG₁κ), and alemtuzumab (Campath^®, Genzyme, Cambridge, MA, USA; anti-CD52 humanized IgG₁κ). Anti-CD3 murine IgG_2a mAb (clone OKT3; BioLegend, San Diego, CA, USA) and anti-CD28 SA murine IgG₁κ mAb (clone ANC28.1/5D10; Ancell Corp., Bayport, MN, USA) were used as reference mAb. Mouse IgG₁ antibody (BioLegend), mouse IgG_2a antibody (BioLegend), human IgG₁ antibody (Eureka Therapeutics, Emeryville, CA, USA), and human IgG₂ antibody (Sigma, St. Louis, MO, USA) were used as isotype controls. Lipopolysaccharide (LPS; from Escherichia coli Type 055:B5) and phytohemagglutinin (PHA) (both Sigma) were used as positive controls.

RPMI-1640 medium, penicillin–streptomycin, fetal bovine serum (FBS), and phosphate-buffered saline (PBS, pH 7.2) were purchased from Thermo Fisher Scientific (Waltham, MA, USA). CTL anti-aggregate wash supplement 20× was obtained from Cellular Technology Limited (Cleveland, OH, USA). The F(ab’)₂ fragments purchased were: anti-human CD64 F(ab’)₂ fragment (clone 10.1), anti-human CD32 F(ab’)₂ fragment (clone 7.3), anti-human CD16 F(ab’)₂ fragment (clone 3G8), and mouse IgG₁ F(ab’)₂ fragment (clone MOPC 31 C) (all Ancell Corporation).

Immobilization of antibodies

Antibody solutions diluted in sterile PBS were added at 5 μl/well and air-dried onto the surface of wells of 96-well polypropylene plates (untreated 96-well U-bottom microtiter plates; Corning Inc., Corning, NY, USA) by overnight incubation at room temperature. The concentrations of antibodies were set at 0.1, 0.3, 1, and 3 μg/well based on preliminary in-house data and previous publications (Findlay et al. 2010, 2011; Wolf et al. 2012; Vessillier et al. 2015). Each coated well was then washed twice with 200 μl sterile PBS. LPS and PHA at a 5 μg/ml final concentration then added to the wells just before the assay without immobilization.

PBMC

Human PBMC were purchased from Cellular Technology Limited. Use of human-derived test materials was approved by the Research Ethics Committee of Daiichi-Sankyo Co., Ltd. (Tokyo, Japan). The requirement for consent was waived by the Ethics Committee.

Cell culture and stimulation

Frozen PBMC from six donors were each thawed at 37 °C and washed with CTL-Thaw Solution (CTL anti-aggregate wash supplement [20×] diluted 20-fold with RPMI-1640 supplemented with 2 mM L-glutamine, 100 U penicillin/ml, and 100 μg streptomycin/ml). The PBMC were then re-suspended in 10% FBS-RPMI1640 (RPMI-1640 with 10% FBS, 2 mM L-glutamine, 100 U penicillin/ml, and 100 μg streptomycin/ml), seeded at 2 × 10⁵ cells in 200 μl volumes into antibody-immobilized wells of 96-well polypropylene plates (see “immobilization of antibodies” in Materials and Methods), and incubated at 37 °C under 5% CO₂ for 24 or 48 hr, in duplicate (non-HDC method). For HDC, 1.5 × 10⁷ PBMC (in 1.5 ml 10% FBS-RPMI-1640) were placed into the wells of a 24-well polystyrene plate (Greiner Bio-One International GmbH, Kremsmünster, Austria) and then incubated at 37 °C under 5% CO₂ for 24 hr to enhance cell reactivity. Culture conditions for the HDC were based on previous publications (Römer et al. 2011; Bartholomaeus et al. 2014; Hussain et al. 2015, 2019). After HDC, the PBMC were inoculated into antibody-immobilized wells as described in the non-HDC method.

For the F_cγR blockade experiments, HDC-PBMC were pretreated with each F(ab’)₂ fragment (at 10 μg/ml) for 45 min at 4 °C before stimulation in antibody-coated wells, in duplicate. Based on preliminary data, the test antibody concentration was set at 3 μg/well, at which the maximum cytokine release responses induced by the antibodies were expected.

Cytokine measurements

At the end of the incubations, culture supernatant from each well was collected after centrifugation (500 × g, room temperature, 10 min). Concentrations of TNFα, IFNγ, IL-2, IL-6, IL-8, and IL-10, and macrophage inflammatory protein (MIP)-1α in each supernatant were then measured using a Milliplex MAP Human Magnetic Cytokine/Chemokine Bead Panel (Merck, Darmstadt, Germany) in a BioPlex^® system (BioPlex 100 array reader with BioPlex Manager software; BioRad, Hercules, CA, USA), according to manufacturer instructions.

Measurement of cell proliferation by WST-8

Three hours prior to the end of incubation, all of the PBMC suspension (i.e., 200 μl) in each well was transferred to a polystyrene flat-bottomed 96-well plate. A total of 20 μl Cell Counting Kit-8 solution containing water-soluble tetrazolium salts-8 (WST-8) (Dojindo Labs, Kumamoto, Japan) was added to each well, and the plate was incubated for 3 hr at 37 °C under 5% CO₂. At the end of incubation, formazan production was assessed by measuring the optical density (OD) at 450 nm (reference wavelength 650 nm) using a SPECTRAmaxPLUS microplate reader with SOFTmaxPRO software (Nihon Molecular Devices Co., Tokyo, Japan).

Statistical analysis

Mean values of maximum responses to 0.1, 0.3, 1, or 3 μg mAb/well for each donor were calculated using Excel software (Office 2013; Microsoft, Redmond, WA, USA) (Findlay et al. 2010, 2011). The maximum responses of mAb were statistically compared to those for panitumumab using a Student’s t-test (homogenous) or Welch’s t-test (not homogenous). In addition, differences in maximum response between HDC and non-HDC conditions were analyzed using these same tests. All data were analyzed using SAS System Release 8.2 software (SAS Institute, Cary, NC, USA). A p-value < 0.05 was considered significant.

In the F_cγR blockade experiments, the percent cytokine release induced by immobilized antibodies from F(ab’)₂ pretreated PBMC relative to that of mouse IgG₁ F(ab’)₂ pretreated PBMC was also calculated. For comparisons of F(ab’)₂ fragment treatment, cytokine concentrations were analyzed using a Student’s (homogeneous) or Welch’s t-test (not homogeneous).

Results

Enhanced response of cytokine release from PBMC by various mAb under HDC conditions

With PBMC from more than half the donors, trastuzumab, rituximab, and alemtuzumab apparently increased (≥ 3-fold) the release of almost all the measured cytokines except for IL-2 under both non-HDC and HDC conditions when compared with effects from PBS. Panitumumab (negative control) also increased release of almost all the cytokines except IL-2 under HDC conditions and except IL-2 and IL-10 under non-HDC conditions (vs. PBS). However, levels of cytokine release induced by panitumumab were clearly lower than those induced by the other three mAb under both non-HDC and HDC conditions (Table 1).

Table 1. Cytokine release from human PBMC induced by various immobilized antibodies.

	TNFα (pg/ml)		IFNγ (pg/ml)		IL-2 (pg/ml)		IL-6 (pg/ml)
Ab	HDC (−)	HDC (+)	HDC (−)	HDC (+)	HDC (−)	HDC (+)	HDC (−)	HDC (+)
Panitumumab	447 ± 243	872 ± 667	10 ± 7	18 ± 17	11 ± 6	9 ± 6	315 ± 361	258 ± 400
Trastuzumab	1701 ± 639^††	2260 ± 939^†	167 ± 259	71 ± 45^§	7 ± 5	8 ± 7	2006 ± 2115	80 ± 135
Rituximab	1475 ± 824^§	1753 ± 889	147 ± 312	52 ± 27^†	6 ± 6	9 ± 6	2410 ± 3203	27 ± 31
Alemtuzumab	1430 ± 780^§	5074 ± 2812^#,§	119 ± 113	299 ± 365	6 ± 7	11 ± 8	2180 ± 1720^§	1171 ± 1606
Anti-human CD3 mAb	4159 ± 1362^§§	10768 ± 3839^##,§§	12736 ± 12549	26551 ± 24321^§	31 ± 14^††	146 ± 153	1075 ± 1083	838 ± 758
Anti-human CD28 SA mAb	13689 ± 9120	32740 ± 12166^*,§§	167691 ± 185393	184273 ± 124381^§§	11681 ± 8146^§	21679 ± 13991^§	3736 ± 2492^§	3065 ± 1526^§§
PBS	35 ± 34	36 ± 21	3 ± 3	6 ± 9	7 ± 5	8 ± 4	77 ± 106	14 ± 23
Human IgG1 Ab	2393 ± 1078	3449 ± 2183	166 ± 292	85 ± 66	7 ± 6	12 ± 10	4159 ± 3017	553 ± 1217*
Human IgG2 Ab	1243 ± 844	932 ± 455	168 ± 273	51 ± 58	13 ± 7	10 ± 7	2260 ± 3046	35 ± 48
Mouse IgG1 Ab	890 ± 724	658 ± 384	99 ± 125	7 ± 9	11 ± 5	9 ± 6	2267 ± 2712	74 ± 151
Mouse IgG2a Ab	1134 ± 734	1513 ± 987	197 ± 299	48 ± 56	13 ± 7	9 ± 6	2651 ± 3324	85 ± 152
LPS	2255 ± 694	9773 ± 4485^##	1004 ± 925	4079 ± 4968	6 ± 3	10 ± 7	6743 ± 2136	18680 ± 4448**
PHA	4302 ± 1531	8520 ± 3081*	14751 ± 11711	46155 ± 32634	522 ± 174	2923 ± 1560^#	6337 ± 2048	8920 ± 3704
	IL-8 (pg/ml)		IL-10 (pg/ml)		MIP-1α (pg/ml)
Ab	HDC (−)	HDC (+)	HDC (−)	HDC (+)	HDC (−)	HDC (+)
Panitumumab	29752 ± 17498	30670 ± 15700	10 ± 7	16 ± 15	4172 ± 2613	7133 ± 4275
Trastuzumab	80787 ± 13971^††	72653 ± 30762^†	21 ± 11	18 ± 11	9609 ± 5471	16884 ± 10123
Rituximab	69528 ± 22765^††	59461 ± 27416^†	22 ± 17	17 ± 11	9521 ± 4693^†	15478 ± 10415
Alemtuzumab	75555 ± 20952^††	132635 ± 54324^*,§§	20 ± 14	49 ± 28^*,†	7458 ± 5163	25748 ± 20217
Anti-human CD3 mAb	52335 ± 23611	77460 ± 36608^§	2689 ± 1123^§§	5675 ± 2824^*,§§	11398 ± 6015^†	24675 ± 11504^*,§
Anti-human CD28 SA mAb	73767 ± 35004^†	94046 ± 30799^††	735 ± 328^§§	1521 ± 610^*,§§	16869 ± 12021^§	24686 ± 11495^§
PBS	2708 ± 1680	2134 ± 839	7 ± 6	4 ± 2	86 ± 67	64 ± 43
Human IgG1 Ab	86839 ± 18289	102142 ± 41129	34 ± 19	21 ± 15	9595 ± 5915	17216 ± 11048
Human IgG2 Ab	54804 ± 31507	41241 ± 12546	22 ± 17	17 ± 11	7350 ± 4724	7672 ± 4127
Mouse IgG1 Ab	51888 ± 38017	29983 ± 14752	25 ± 19	12 ± 6	5066 ± 4007	4760 ± 3753
Mouse IgG2a Ab	54219 ± 32924	51568 ± 19417	25 ± 17	15 ± 10	6006 ± 3519	9108 ± 5302
LPS	35620 ± 13987	152723 ± 22079**	662 ± 364	1272 ± 669	7441 ± 4184	14476 ± 5866*
PHA	30283 ± 15867	114862 ± 32006**	1524 ± 597	2255 ± 1275	10710 ± 5130	19517 ± 10637

Ab: antibody, mAb: monoclonal antibody, HDC: high-density preculture, LPS: lipopolysaccharide, PHA: phytohemagglutinin, SA: super-agonist. A 0.2 ml suspension of human PBMC (2 × 105 cells/well) was incubated with immobilized Ab (0.1, 0.3, 1, and 3 µg/well) for 48 hr exposure. Cytokine levels in well supernatants were measured after the 48 hr exposure. Maximum responses to mAb are shown. Values shown are the means ± SD of six donors from six independent experiments. *p < 0.05 (Student's t-test), **p < 0.01 (Student's t-test), ^#p < 0.05 (Welch's test), ##p < 0.01 (Welch's test): compared with the response in HDC (-). ^†p < 0.05 (Student's t-test), ^††p < 0.01 (Student's t-test), ^§p < 0.05 (Welch's test), ^§§p < 0.01 (Welch's test) compared with the response of panitumumab in the corresponding conditions.

TNFα, IL-8, and IL-10 release due to alemtuzumab under HDC conditions were significantly higher vs. those under non-HDC conditions. In contrast, neither trastuzumab, rituximab, nor panitumumab caused a difference in cytokine release levels between conditions (Table 1). In addition, when compared with panitumumab, all cytokines except IL-2 showed a statistically significant increase or apparent (≥ 3-fold) increase in more than half the donor PBMC tested by alemtuzumab under HDC conditions; significant or apparent increases were seen only for TNFα, IFNγ, IL-6, and IL-8 under non-HDC conditions. On the other hand, no increases were found in the number of increased cytokines induced by rituximab and trastuzumab under HDC conditions when compared with that under non-HDC conditions. Specifically, cytokines that gave rise to a statistically-significant increase or apparent (≥ 3-fold) increase in more than half the donor PBMC tested with rituximab under HDC conditions were TNFα, IFNγ, and IL-8; under non-HDC conditions, the cytokines displaying such outcomes were TNFα, IFNγ, IL-6, IL-8, and MIP-1α. With trastuzumab, under HDC conditions, the cytokines that displayed a statistically significant or apparent (≥ 3-fold) increase in more than half the donor PBMC tested were TNFα, IFNγ, IL-8, and MIP-1α; under non-HDC conditions, TNFα, IFNγ, IL-6, IL-8, and IL-10 displayed these outcomes (Tables 1 and 2). As for other changes, there was a tendency to lower IL-6 release under HDC conditions as a result of PBMC treatment with panitumumab, trastuzumab, rituximab, or alemtuzumab exposure compared with the levels noted under non-HDC conditions (Table 1).

Table 2. Cytokine release response to various immobilized antibodies by donor PBMC.

	TNFα (pg/ml)				IFNγ (pg/ml)
	HDC (−)		HDC (+)		HDC (−)		HDC (+)
Ab	Fold-change range	# responding donors	Fold-change range	# responding donors	Fold-change range	# responding donors	Fold-change range	# responding donors
Trastuzumab	2.3–6.6	5	1.1–7.7	4	2.3–35.7	5	1.8–8.9	5
Rituximab	2.0–4.7	3	0.7–4.5	3	1.4–40.7	4	1.7–10.4	2
Alemtuzumab	1.3–5.1	4	1.7–11.2	5	1.5–51.6	4	3.8–27.4	6
Anti-human CD3 mAb	5.6–18.0	6	5.0–36.0	6	282.0–3833.1	6	664.4–3304.4	6
Anti-human CD28 SA mAb	16.3–55.6	6	16.9–101.9	6	3472.7–60752.6	6	3511.2–26743.7	6
	IL-2 (pg/ml)				IL-6 (pg/ml)
	HDC (−)		HDC (+)		HDC (−)		HDC (+)
Ab	Fold-change range	# responding donors	Fold-change range	# responding donors	Fold-change range	# responding donors	Fold-change range	# responding donors
Trastuzumab	0.2–1.1	0	0.0–4.3	1	3.0–11.4	5	0.0–12.2	1
Rituximab	0.0–1.3	0	0.0–4.6	1	2.1–20.6	3	0.0–4.9	1
Alemtuzumab	0.0–1.1	0	0.0–5.0	1	3.7–14.8	6	0.1–483.6	5
Anti-human CD3 mAb	1.5–8.2	2	1.9–48.9	5	2.8–6.1	5	0.1–475.1	5
Anti-human CD28 SA mAb	424.8–1399.0	6	843.8–4737.0	6	6.9–37.3	6	2.3–1208.9	5
	IL-8 (pg/ml)				IL-10 (pg/ml)
	HDC (−)		HDC (+)		HDC (−)		HDC (+)
Ab	Fold-change range	# responding donors	Fold-change range	# responding donors	Fold-change range	# responding donors	Fold-change range	# responding donors
Trastuzumab	1.8–5.0	3	1.4–7.0	2	0.8–9.8	3	0.0–6.4	1
Rituximab	1.7–4.7	2	1.0–3.9	1	0.4–12.9	1	0.0–8.5	1
Alemtuzumab	1.4–4.8	3	2.2–8.8	4	0.3–6.4	2	0.4–11.4	4
Anti-human CD3 mAb	1.3–2.6	0	1.4–6.6	2	145.4–689.3	6	103.1–1181.4	6
Anti-human CD28 SA mAb	1.7–4.1	1	1.9–5.8	3	30.5–272.6	6	33.4–415.6	6
	MIP-1α (pg/ml)
	HDC (−)		HDC (+)
Ab	Fold-change range	# responding donors	Fold-change range	# responding donors
Trastuzumab	0.9–4.0	2	1.0–3.9	3
Rituximab	1.6–4.2	1	0.8–3.5	1
Alemtuzumab	0.9–2.2	0	1.6–7.0	3
Anti-human CD3 mAb	1.9–4.4	2	1.9–9.1	4
Anti-human CD28 SA mAb	2.3–4.6	5	2.4–8.0	6

Ab: antibody, mAb: monoclonal antibody, HDC: high-density preculture, SA: super-agonist.

Fold-change in cytokine level associated with each mAb relative to corresponding levels induced by panitumumab. Fold-change range shows minimum-maximum fold-change of cytokines for all six donors. Number of responding donors is number of positive responding donors showing a fold-change of ≥3. Results in which more than half of the donors showed a positive response are shown in bold.

Compared with PBS, anti-CD3 mAb induced significant increases in release of all the cytokines measured under both non-HDC and HDC conditions. There was also a significant increase in TNFα, IL-10, and MIP-1α release caused by anti-CD3 mAb under HDC conditions compared with levels under non-HDC conditions (Table 1). Similarly, anti-CD28 SA mAb induced significant increases in release of all measured cytokines compared with that due to PBS under non-HDC and HDC conditions. Lastly, compared to findings under non-HDC conditions, there were significant increases in TNFα and IL-10 release induced by anti-CD28 SA mAb under HDC conditions.

Four isotype controls (human IgG₁ and IgG₂, mouse IgG₁ and IgG_2a) also caused increased release of almost all the measured cytokines (except IL-2) compared with PBS under both non-HDC and HDC conditions. As with the other mAb tested, there was a tendency to lower IL-6 release under HDC conditions compared with under non-HDC conditions (Table 1). Both LPS- and PHA-positive controls induced marked cytokine release under both non-HDC and HDC conditions, with the increase higher under HDC vs. non-HDC conditions.

For Readers interested in examining cytokine release at the level of each individual donor, these data are provided here in a Supplementary file.

Cell proliferation in PBMC by mAb under HDC conditions

Anti-CD3 mAb and anti-CD28 SA mAb increased cell proliferation under both non-HDC and HDC conditions. Furthermore, the proliferative response due to anti-CD3 mAb was higher under HDC than under non-HDC conditions (Table 3). Panitumumab, trastuzumab, rituximab, and alemtuzumab did not induce cellular growth in either condition (Table 3).

Table 3. Cell proliferation induced in donor PBMC by various immobilized antibodies.

	Cell proliferation (OD 450)
Ab	HDC (−)	HDC (+)
Panitumumab	0.366 ± 0.027	0.364 ± 0.052
Trastuzumab	0.344 ± 0.036	0.363 ± 0.032
Rituximab	0.371 ± 0.056	0.382 ± 0.050
Alemtuzumab	0.368 ± 0.066	0.399 ± 0.047
Anti-human CD3 mAb	0.870 ± 0.276	1.406 ± 0.475*
Anti-human CD28 SA mAb	1.046 ± 0.347	1.264 ± 0.407
PBS	0.387 ± 0.046	0.414 ± 0.036
Human IgG1 Ab	0.344 ± 0.036	0.375 ± 0.039
Human IgG2 Ab	0.372 ± 0.043	0.353 ± 0.030
Mouse IgG1 Ab	0.370 ± 0.050	0.370 ± 0.036
Mouse IgG2a Ab	0.379 ± 0.063	0.359 ± 0.025
LPS	0.323 ± 0.028	0.501 ± 0.057**
PHA	0.746 ± 0.155	1.151 ± 0.339*

Ab: antibody, mAb: monoclonal antibody, HDC: high-density pre-culture, LPS: lipopolysaccharide, PHA: phytohemagglutinin, SA: super-agonist.

A 0.2 ml suspension of human PBMC (2 × 10⁵ cells/well) was incubated with immobilized antibodies (0.1, 0.3, 1, and 3 μg/well) for 48 hr. Cell proliferation was determined using a WST-8 assay over the final 3 hr of incubation. Maximum responses to mAb/Ab are shown. Values shown are means ± SD of six donors of six independent experiments. *p < 0.05 (Student’s t-test), **p < 0.01 (Student’s t-test) compared with that in HDC (−).

Time course of cytokine release from/cell proliferation in PBMC induced by various mAb

Almost all the releases of cytokines and cell proliferation induced by anti-CD3 and anti-CD28 SA mAb under HDC conditions were time-dependent and more pronounced at 48 than at 24 hr. The results for the release of TNFα (as a representative cytokine) are shown in Figure 1. In contrast, the release of cytokines due to panitumumab, trastuzumab, rituximab, and alemtuzumab was not affected by the length of the incubation period (Figure 1).

Figure 1. TNFα release and cell proliferation induced by various immobilized antibodies from human peripheral blood mononuclear cells for 24 or 48 hr. mAb: monoclonal antibody; SA: super-agonist. A 0.2 ml suspension of human PBMC (2 × 10⁵ cells/well) following high-density pre-culture (10⁷ cells/ml) was incubated with immobilized Ab (0.1, 0.3, 1, and 3 μg/well) for 24 or 48 hr. (A) TNFα levels in the supernatants after 24 or 48 hr of exposure. (B) Cell proliferation determined by WST-8 assay over last 3 hr of 24 or 48 hr incubations. Values shown are means ± SD of 5–6 donors from six independent experiments. *p < 0.05, **p < 0.01 (Student’s t-test), ^#p < 0.05, ^##p < 0.01 (Welch’s test): compared with values at 24 hr.

Inhibition of cytokine release by F_cγR blockade

Cytokine release induced by alemtuzumab was reduced by pretreatment with anti-F_cγRI, -F_cγRII, and -F_cγRIII F(ab’)₂ fragments. Further, the suppressive effect of anti-F_cγRIII F(ab’)₂ on alemtuzumab-induced cytokine release was more pronounced than that of anti-F_cγRI and anti-F_cγRII (Table 4). The release of multiple cytokines due to trastuzumab was also suppressed by F_cγRIII blockade, but not affected by F_cγRI or F_cγRII. Interestingly, though release of cytokines induced by rituximab was also suppressed most by F_cγRIII blockade, IL-10 release by rituximab was only suppressed by F_cγRII blockade. For the release of cytokines caused by panitumumab, the IL-6 release was only suppressed by F_cγRIII blockade. The cytokine release due to isotype control antibodies was also reduced by F_cγRI, F_cγRII, or F_cγRIII blockade. Release of cytokines due to LPS and PHA was not affected by the inhibition of the F_cγR (data not shown).

Table 4. Cytokine release induced by various immobilized antibodies from F(ab’)₂ fragment pretreated human PBMC.

mAb/Ab	F(ab')2 fragment	TNFα		IFNγ		IL-2		IL-6		IL-8		IL-10		MIP-1α
		(pg/ml)	(%)^a	(pg/ml)	(%)	(pg/ml)	(%)	(pg/ml)	(%)	(pg/ml)	(%)	(pg/ml)	(%)	(pg/ml)	(%)
Panitumumab	Mouse IgG1	834 ± 358	–	3 ± 2	–	6 ± 4	–	13 ± 9	–	33126 ± 10150	–	2 ± 1	–	12146 ± 6803	–
	Anti-FcγRI	627 ± 176	75	2 ± 1	91	8 ± 5	119	46 ± 72	338	26905 ± 6841	81	2 ± 1	109	10104 ± 6909	83
	Anti-FcγRII	949 ± 650	114	14 ± 28	540	8 ± 4	121	138 ± 301	1021	34710 ± 11441	105	4 ± 5	201	10618 ± 5179	87
	Anti-FcγRIII	496 ± 262	59	2 ± 2	78	5 ± 3	79	9 ± 4	63^#	22262 ± 10439	67	2 ± 1	89	7673 ± 6006	63
Trastuzumab	Mouse IgG1	1585 ± 569	–	80 ± 81	–	6 ± 4	–	65 ± 59	–	54744 ± 24372	–	3 ± 3	–	14369 ± 10801	–
	Anti-FcγRI	1756 ± 559	111	89 ± 86	112	5 ± 3	80	191 ± 195	295	51732 ± 11369	94	3 ± 2	93	13240 ± 8643	92
	Anti-FcγRII	1541 ± 348	97	100 ± 111	125	7 ± 6	123	79 ± 110	123	52918 ± 20553	97	2 ± 1	67	12329 ± 7693	86
	Anti-FcγRIII	1012 ± 591	64^#	28 ± 35	35^##	4 ± 2	68	22 ± 14	34^#	33792 ± 17113	62	1 ± 2	36	7552 ± 3771	53
Rituximab	Mouse IgG1	1299 ± 717	–	38 ± 50	–	4 ± 3	–	24 ± 15	–	46230 ± 23286	–	2 ± 1	–	11188 ± 7998	–
	Anti-FcγRI	1386 ± 527	107	39 ± 49	103	3 ± 3	75	52 ± 54	211	46840 ± 15503	101	1 ± 1	57	12657 ± 7732	113
	Anti-FcγRII	1117 ± 396	86	33 ± 35	87	4 ± 2	88	32 ± 20	132	43291 ± 11688	94	1 ± 1	62^#	9599 ± 4653	86
	Anti-FcγRIII	756 ± 525	58^#	9 ± 8	24^##	3 ± 3	69^#	9 ± 3	37^#	27798 ± 15283	60^#	1 ± 1	54	5820 ± 3266	52
Alemtuzumab	Mouse IgG1	4090 ± 2258	–	298 ± 389	–	3 ± 2	–	1116 ± 1550	–	124309 ± 45291	–	42 ± 34	–	20157 ± 13475	–
	Anti-FcγRI	2198 ± 801	54^#	200 ± 205	67	3 ± 2	86	311 ± 243	28	68553 ± 19565*	55^#	24 ± 25	58	14255 ± 6362	71
	Anti-FcγRII	2427 ± 910	59^#	269 ± 316	90	3 ± 2	91	364 ± 211	33	90795 ± 28831	73	34 ± 35	81	18938 ± 14971	94
	Anti-FcγRIII	1239 ± 692^#	30^##	69 ± 54	23^##	2 ± 1	67^#	109 ± 84	10^##	40699 ± 19697**	33^##	16 ± 17	39^#	8828 ± 3334	44
Human IgG1 Ab	Mouse IgG1	3257 ± 2367	–	83 ± 156	–	6 ± 5	–	367 ± 427	–	76234 ± 22876	–	12 ± 16	–	17838 ± 11449	–
	Anti-FcγRI	1744 ± 756	54^#	88 ± 129	105	5 ± 6	86	171 ± 181	47	51125 ± 14977*	67^#	4 ± 3	35	11787 ± 7302	66^#
	Anti-FcγRII	2446 ± 1122	75^#	90 ± 129	108	15 ± 8*	227	312 ± 345	85	75232 ± 20096	99	6 ± 4	50	15874 ± 10765	89
	Anti-FcγRIII	1826 ± 1168	56^##	18 ± 13	21	12 ± 9	181	204 ± 326	55^#	50841 ± 22035	67^##	5 ± 5	40^#	11682 ± 7503	65^##
Human IgG2 Ab	Mouse IgG1	1142 ± 622	–	6 ± 3	–	7 ± 2	–	84 ± 163	–	42019 ± 15412	–	3 ± 1	–	12545 ± 10775	–
	Anti-FcγRI	946 ± 574	83	8 ± 8	131	7 ± 6	108	32 ± 25	38	34973 ± 14853	83	2 ± 2	85	8336 ± 4727	66
	Anti-FcγRII	969 ± 401	85	7 ± 7	124	6 ± 4	94	25 ± 13	29	40061 ± 12630	95	2 ± 2	77	10534 ± 6447	84
	Anti-FcγRIII	647 ± 413	57^##	4 ± 4	73	5 ± 5	82	10 ± 7	12^##	27174 ± 11740	65^##	2 ± 1	60^#	6635 ± 3442	53^#

Ab: antibody, LPS: lipopolysaccharide, PHA: phytohemagglutinin.

High-density precultured human PBMC (2 × 10⁵ cells/well) were incubated with F(ab’)₂ fragments of neutralizing antibodies to F_cγRI (CD64), F_cγRII (CD32), or F_cγRIII (CD16) for 45 min on ice. Then, a 0.2 ml suspension was incubated with immobilized antibodies (3 μg/well) for 48 hr, after which cytokine levels in well supernatants were measured. Values shown are means ± SD of six donors from two independent experiments.

^aPercentage cytokine release induced by various immobilized antibodies from F(ab’)₂ fragment pretreated PBMC compared with that due to mouse IgG₁ F(ab’)₂ fragment treatment is shown. Percentage = 100 × [absolute average value of six donors in each F(ab’)₂ treatment/absolute average value of six donors with mouse IgG₁ F(ab’)₂ treatment]. *p < 0.05 (Student’s t-test), **p < 0.01 (Student’s t-test), ^#p < 0.05 (Welch’s test), ^##p < 0.01 (Welch’s test) compared with response with mouse IgG₁ F(ab’)₂ fragment treatment.

Discussion

In this study, cytokine release due to trastuzumab, rituximab, or alemtuzumab in the air-drying solid-phase method was higher than that due to panitumumab under both non-HDC and HDC conditions. Further, unlike for trastuzumab, rituximab, and panitumumab, the cytokine release caused by alemtuzumab was more pronounced under HDC conditions. Consistent with the levels of released cytokines under HDC conditions in the present study, the incidence of infusion-related reaction (IRR; a disorder characterized by adverse reaction [such as chills, fever, nausea, headache, etc.] that typically develops during or several hours after the infusion) in a clinical context was reported to be 40% for trastuzumab (Genentech 2018), 77% for rituximab (Genentech 2019), and 97% for alemtuzumab (Sanofi 2015), whereas that for panitumumab was reported to be very low (4%) (Chung 2008). Therefore, the new CRA system developed here that incorporated HDC into an air-drying solid-phase CRA could detect the potential risk of CRS associated with the various types of mAb that induce cytokine release via F_cγR in humans.

Here, alemtuzumab caused more pronounced release of TNFα, IL-8, and IL-10 from PBMC under HDC conditions than that under non-HDC conditions. It has been suggested that the release of TNFα, IFNγ, and IL-6 into the blood circulation that is associated with alemtuzumab originates from NK cells, monocytes, and neutrophils (Wing et al. 1996; Hu et al. 2009; Siders et al. 2010; Stebbings et al. 2013). In addition, Hussain et al. (2015, 2019) reported that expression levels of F_cγRIIb and F_cγRIIIa on monocytes increased under HDC conditions, but those of F_cγRI or F_cγRIIa on monocytes under these conditions only slightly increased or remained unchanged, respectively. Further, these authors reported that expression of F_cγRIIIa on NK cells under HDC were comparable to that under non-HDC conditions. It was also reported that immobilized anti-F_cγRIII antibody induced cellular activation accompanied by TNFα and IFNγ production from NK cells (Romee et al. 2013). Moreover, monocytes expressing F_cγRIII were reported to show ADCC activity depending on the affinity to F_cγRIII, similar to the case for NK cells (Yeap et al. 2016), and monocytes were further reported to release TNFα by crosslinking to F_cγRIII (Abrahams et al. 2000; Kramer et al. 2004).

Taking together this information and the present results showing that alemtuzumab-induced release of cytokines under HDC conditions was strongly inhibited by F_cγRIII blockade, it was suggested that alemtuzumab induced cytokine release from NK cells and monocytes, and that the levels of release under HDC conditions were more pronounced due to the contributions from cytokines that were being released from monocytes with increased F_cγRIIIa expression (due to HDC). Similarly, release of cytokines caused by trastuzumab and rituximab was suppressed by F_cγRIII blockade, indicating these mAb also cause release of cytokines via F_cγRIII under HDC conditions. However, unlike alemtuzumab, neither trastuzumab nor rituximab enhanced cytokine release under HDC conditions, implying that increased F_cγRIII expression on monocytes and cytokine release from monocytes might not be mainly attributable to the cytokine release induced by these mAb under HDC conditions.

T-cell agonists anti-CD3 and anti-CD28 SA mAb induced significant release of cytokines and cell proliferation, especially under HDC conditions. Similar cytokine release and cell proliferation were reported in CRA under HDC conditions for TGN1412, another anti-CD28 SA mAb, or OKT3 (Römer et al. 2011; Bartholomaeus et al. 2014; Hussain et al. 2015). Römer et al. (2011) suggested that co-localization of CD3 and phosphorylated tyrosine on T-cells increased T-cell reactivity under HDC conditions. Lühder et al. (2003) proposed that anti-CD28 SA mAb, unlike conventional anti-CD28 mAb, recognized the proximal epitope of the CD28 antigen (capable of oligomerizing CD28 on T-cell surface), resulting in T-cell receptor (TCR)- independent T-cell activation. It was also postulated that increased F_cγRIIb expression on monocytes under HDC conditions (Hussain et al. 2015, 2019) caused clustering of CD28 on the T-cell surface, leading to T-cell activation. This would be along the lines of interactions between CD80/CD86 on antigen-presenting cells and CD28 on T-cells.

In the present study, compared with PBS, panitumumab associated with a low risk of CRS caused mild increases in cytokine release in both HDC and non-HDC conditions. In solid-phase CRA, it is well known that antibodies induce nonspecific cytokine release via the binding of their F_c section to effector cell F_cγR (Findlay et al. 2011; Eastwood et al. 2013; Vessillier et al. 2015; Grimaldi et al. 2016). Cytokine release by panitumumab was inhibited by F_cγRIII blockade under HDC conditions in this study, suggesting this class of cytokine release was an F_cγR- dependent nonspecific effect. In addition, the order of intensity of cytokine release caused by the mAb tested was “trastuzumab (humanized IgG₁) ≈ rituximab (humanized IgG₁) > panitumumab (human IgG₂), and human IgG₁ isotype control antibody > human IgG₂ isotype control antibody” under both HDC and non-HDC conditions. Moreover, the degree of inhibition by F_cγRIII blockade was small for panitumumab (humanized IgG₂) compared with the findings for alemtuzumab, rituximab, or trastuzumab (each a humanized IgG₁). Taking into account that the affinity of human IgG₁ for F_cγR is higher than that of human IgG₂ for all of F_cγRI, F_cγRII, and F_cγRIII (Brennan et al. 2010), the intensity of cytokine release and the effect of F_cγR blockade in this study might be related to the affinity of each isotype to F_cγR as described above.

In conclusion, it would seem that incorporating HDC into air-drying solid-phase CRA could sensitively capture cytokine-releasing effects induced via by the binding of the F_c region of mAb (such as alemtuzumab) with F_cγR on immune cells.

Acknowledgments

The authors thank Hitomi Terada for excellent support in the experiments.

Disclosure statement

The authors declare no conflicts of interest. The authors alone are responsible for the content of this paper.