Itacitinib (INCB039110), a JAK1 Inhibitor, Reduces Cytokines Associated with Cytokine Release Syndrome Induced by CAR T-cell Therapy

This article is featured in Highlights of This Issue, p. 6075

T cells engineered to express a chimeric antigen receptor (CAR) have recently been approved by the FDA, and commercially available CAR T-cell therapies (axicabtagene ciloleucel and tisagenlecleucel) have resulted in an overall response rate of almost 80% (1–4), including long-term relapse-free survival in patients with B-cell leukemia and lymphoma (5). However, an exaggerated immune response by the adoptively transferred CAR T cells, as well as the host immune system, can lead to a severe cytokine release syndrome (CRS) in up to 40% of treated patients (6, 7). CRS manifests as a rapid immune reaction driven by a massive release of inflammatory cytokines. While individual cytokine levels differs between patients, severe CRS is generally very similar to that seen in hemophagocytic lymphohistiocytosis (HLH) or macrophages activation syndrome (MAS; ref. 8), with elevation of both canonical T-cell cytokines (i.e., IFNγ) as well as cytokines compatible with HLH/MAS (i.e., IL6 and IL10; ref. 9). Typified by chills and fevers, CRS can also progress to a more severe, even life-threatening reaction (10, 11).

To date, low-grade CRS is treated symptomatically with antipyretics and fluids (7). Grade > 2 CRS can be treated with tocilizumab (an anti-IL6 receptor), recently approved by the FDA and the European Medicines Agency for treatment of severe and life-threatening CRS (12, 13). Importantly, therapeutic administration of tocilizumab does not influence peak CAR T-cell numbers nor clinical outcomes (14). However, treatment with corticosteroids is also needed in the case of tocilizumab-refractory patients, and concerns remain over the effect of high-dose corticosteroids on CAR T-cell expansion and antitumor activity (14). To date, there are no preventative strategies, and we rely on therapeutic treatments. Thus, the ability to prophylactically inhibit hyperinflammatory states and the incidence of CRS would represent a new clinical management strategy for patients receiving CAR T therapy.

JAKs play an important role in signal transduction following cytokine and growth factor binding to their cognate receptors. Once activated, JAKs phosphorylate STAT proteins, which results in dimerization and translocation of STATs to the nucleus to activate or suppress gene transcription (15). Aberrant production of JAK-mediated cytokines, such as IL6 and growth factors, has been associated with a number of chronic inflammatory conditions (16, 17). Thus, we reasoned that the blockade of the JAK signaling pathway would attenuate the excessive cytokine response that occurs during CRS without negatively impacting CAR T-cell function. Itacitinib (INCB039110) is a small-molecule inhibitor of the JAK family of protein tyrosine kinases (18). As opposed to other JAK inhibitors developed by Incyte (i.e., ruxolitinib) as well as others (i.e., tofacitanib), itacitinib is a JAK1 selective inhibitor, and is therefore expected to minimize the risk of long-term infection or other complications associated with pan-JAK inhibitors (19), while reducing the levels of cytokines that signal trough JAK1, such as IFNγ, IL6, IL12, or TNFα. In this study, we show that in both in vitro and in vivo preclinical models, itacitinib doses relevant to cellular IC₅₀ (50–100 nmol/L) were more effective than tocilizumab in reducing CRS-related cytokines produced by CD19-CAR T cells. Moreover, itacitinib did not significantly inhibit in vitro proliferation or antitumor killing capacity of CAR T cells, nor, in a lymphoma model, did it affect the antitumor activity of CAR T cells in a prophylactic setting. Together, these data suggest that itacitinib has potential as a prophylactic agent for the prevention of CAR T cell–induced CRS, and a phase II clinical trial of itacitinib for the prevention of CRS induced by CAR T cells is ongoing (NCT04071366).

C57BL/6 and BALB/c animals were purchased from Taconic Biosciences and were approximately 8 weeks old at the time experiments were initiated. OT-1 TCR-transgenic mice and severely immunodeficient NOD-scid IL2R gamma null (NSG) animals were purchased from the Jackson Laboratory. All mice were used in protocols approved by the Institutional Animal Care and Use Committee.

Peripheral blood mononuclear cells (PBMC) were prepared from human whole blood samples using a Ficoll-Hypaque separation method, and T cells were then obtained from the PBMCs by centrifugal elutriation. T cells were maintained in RPMI1640 medium (Thermo Fisher Scientific) supplemented with 10% FBS, 1% HEPES, 2 mmol/L l-glutamine, 0.05 mmol/L 2-mercaptoethanol and 100 μg/mL streptomycin, and 100 units/mL penicillin [complete RPMI or complete media (CM)]. T cells were activated with Dynabeads (Thermo Fisher Scientific; immobilized agonist antibodies against CD3/CD28) at a 3:1 ratio, resuspended at a density of 0.5 × 10⁶ cells/mL in 24-well plates and treated with itacitinib at various concentrations (from 50 to 1,000 nmol/L). The plates were incubated at 37°C in 5% CO₂ atmosphere for 10 days, and the proliferation was determined every other day by bead-based methods (CountBright Absolute Counting Beads, Molecular Probes, Thermo Fisher Scientific). Cultures were replenished every other day with fresh CM.

Luciferase expressing SY5Y neuroblastoma cells (GD-2⁺) were plated in a 96-well plate at 50,000 cells per well. Twenty-four hours later, 150,000 CAR T cells were added to corresponding wells in a final volume of 200 μL. Target cells alone were seeded in parallel to quantify the maximum luciferase expression (relative luminescent units; RLUmax). Seventeen hours later, 100 μL of luciferin substrate (Bright-Glo Luciferase Assay System, Promega) was added to the coculture. Luminescence was measured after a 10-minute incubation using an EnVision (PerkinElmer) plate reader. The percent cell lysis was obtained using the following calculation: [1 − (RLUexperimental)/(RLUmax)] × 100. The experiment was performed in triplicates.

CRS was induced in BALB/c animals by intravenous injection of Concanavalin-A (ConA; 20 mg/kg) or 100 μg of an anti-CD3ϵ antibody (clone 145–2C11). Corresponding animals were orally dosed with 60 or 120 mg/kg of itacitinib 60 minutes prior to CRS induction (prophylactic) or 30 minutes after (therapeutic) CRS induction. Two hours after ConA or anti-CD3 injection, mice were sacrificed and blood was collected into K₂EDTA tubes for cytokine measurement.

Spleens were collected from 6- to 8-week-old C57BL/6 female mice and mechanically dissociated. Splenocytes were cultured in RPMI medium supplemented with 10% FBS and macrophage colony-stimulating factor. On day 6, cells were treated with itacitinib and then treated with 5 ng/mL lipopolysaccharide (LPS) on day 7. The following day, supernatant was collected and cytokines were measured as described below. Six- to 8-week-old female C57BL/6 mice were prophylactically orally dosed with vehicle, or 60 or 120 mg/kg of itacitinib twice a day for 3 days. Mice then received intraperitoneal injections of LPS (5 μg/animal). Two hours after injection, mice were euthanized and 3 mL of sterile saline was injected into the peritoneum to perform a peritoneal lavage. Subsequent cytokine measurements were performed as described below.

Plasma cytokines were measured using multispot assay system, proinflammatory panel 1 [for IFNγ, IL1β, IL2, IL4, IL5, IL6, KC/GRO, IL10, IL1p70, and TNFα; Meso Scale Discovery, Meso Scale Diagnostics (MSD)], following manufacturer's instructions. Briefly, samples, standards, and controls were added at 25 μL per well. The plate was sealed and incubated for 2 hours at room temperature on an orbital shaker (600 rpm). At the end of the incubation, the wells were washed three times using 200 μL PBS + 0.05% Tween 20, soaking for 30 seconds and then discarding. Detection antibody was added at 25 μL per well, and the plate sealed and incubated for 1 hour at room temperature on an orbital shaker (600 rpm). At the end of the incubation, the plate was washed three times as done previously. A total of 150 μL of the MSD Read Buffer was added to each well, and the MSD plates were measured on an MSD Sector S 600 plate reader. The raw data were measured as an electrochemiluminescence signal detected by photodetectors and analyzed using the MSD Discovery Workbench software. A four-parameter logistic fit curve was generated for each analyte using the standards, and the concentration of each sample was calculated.

The OT-1 strain is transgenic for the TCR VαVβ5 specific for the OVA_257–264 peptide (SIINFEKL) and restricted to H2-Kb. Naïve OT-1 CD8 cells were isolated from OT-1 spleens by negative selection using Naïve CD8a⁺ T-Cell Isolation Kit (Miltenyi Biotec).

To measure OT-1 CD8⁺ cell expansion, cells were carboxyfluorescein succinimidyl ester (CFSE)-labeled (2 μmol/L) and resuspended at 5 × 10⁶ cells/mL in complete RPMI, 20 IU IL2, 2 μg/mL of SIINFEKL peptide. and increasing itacitinib concentrations, from 0 to 1,000 nmol/L. Three to five days later, OT-1 cells were collected, stained with an APC-anti-CD8 antibody, and the number of divisions was calculated by measuring the relative CFSE fluorescence intensity of the different conditions by flow cytometry (20).

To study the effect of itacitinib on the antitumor activity of CD8 OT-1 cells, C57BL/6 animals received subcutaneous injections of 0.5 × 10⁶ OVA-expressing EG7 tumor cells into the shaved right flank. At day 7, corresponding animals were orally dosed with itacitinib twice a day at 60 or 120 mg/kg, and at day 10, corresponding animals received an intravenous (tail vein) injection of 5 × 10⁶ naïve OT-1 CD8 cells. Tumor sizes were measured on two perpendicular axes using a digital caliper at least three times a week. Tumor volumes were calculated with the formula:

Where V is tumor volume, W is tumor width, and L is tumor length.

Cell proliferation was measured using CellTiter-Glo Luminescent Cell Viability Assay (Promega), following manufacturer's instructions. Briefly, 1,000 EG7 cells were seeded in a total volume of 200 μL of RPMI + 10% FBS per well in a 96-well plate. The next day, compound plates were reconstituted in complete media with a 10-point dose curve. Cell media of the plate was aspirated and compound with a total volume of 100 μL was incubated with the cells for 72 hours. A total of 100 μL of CellTiter-Glo reagent was added to each well and protected from the light with a plate seal. Plates were shaken at room temperature for 2 minutes to induce cell lysis and then allowed to sit for approximately 10 minutes. The raw data were measured on a TopCount Luminescence Counter (Perkin Elmer) and analyzed in Prism using an inhibitor versus response–variable slope (four parameters) equation.

The CD19-BBζ CAR consisting of a CD8 hinge, 4-1BB costimulatory domain, and CD3ζ signaling domain was generated as described previously (21). PBMCs were stimulated with Dynabeads (3:1 ratio) and then plated in 24-well plates in X-Vivo medium (Lonza) containing IL2 (200 IU/mL), IL7 (10 ng/mL), IL15 (5 ng/mL), and IL21 (5 ng/mL) at 0.5 × 10⁶ cells/mL. Three days later, T cells were transduced using a lentiviral vector expressing the CAR construct, as described elsewhere (22).

Generation of the GD2-E101K CAR lentiviral plasmid was described previously (23). The CAR contains the 14G2a single-chain variable fragment (scFv), as well as an EF1α promoter, CD8 hinge, 4-1BB costimulatory domain, and CD3ζ signaling domain. The anti-EGFR CAR, with 3C10 scFv, was described previously (24). The CAR contains the 3C10 scFV, as well as an EF1α promoter, CD8 hinge, 4-1BB costimulatory domain, and CD3ζ signaling domain.

To study whether JAK1 inhibition affects the antitumor effect of CAR T cells, a xenograft model was used as reported previously (25). Briefly, 6- to 10-week-old immunodeficient NOD-SCID γ_c^−/− (NSG) mice were injected intravenously via tail vein with 2.5 × 10⁶ luciferase-expressing Nalm6 acute lymphoblastic leukemia cells. Corresponding animals received 120 mg/kg of itacitinib per 10 days (orally, twice daily), starting 1 day after the tumor challenge. Mice were then randomized into groups for adoptive transfer of 3 × 10⁶ CAR T cells, nontransduced cells, or vehicle intravenously at day 4. Anesthetized mice were imaged using a Xenogen IVIS Spectrum system (Caliper Life Sciences) once a week. Mice were given an intraperitoneal injection of d-luciferin (150 mg/kg; Caliper Life Sciences). Total flux was quantified using Living Image 4.4 (PerkinElmer) by drawing rectangles of identical area around mice, reaching from head to 50% of the tail length. Background bioluminescence was subtracted for each image individually. Animals were monitored daily, and survival rates were compared between groups.

Six- to 8-week-old male NSG mice were subcutaneously inoculated with luciferase-expressing CD19⁺ NAMALWA lymphoma cells (5 × 10⁶). Mice were weighed and monitored two times a week, and bioluminescence signals were measured every 5 days. At day 5 after tumor challenge, animals started a twice-daily oral treatment of vehicle or INCB039110 (120 mg/kg) for 8 days. At day 8 after tumor challenge, animals received an adoptive transfer of 5 × 10⁶ CD19-CAR T cells. Bioluminiscence was measured as done previously. Animals were monitored daily, and survival rates were compared between groups.

Data are reported as mean ± SD or mean ± SEM in the relevant figures. Differences between two groups were analyzed by nonparametric Mann–Whitney test. Statistical analysis for multiple groups was performed by Kruskal–Wallis with Dunn post hoc test for nonparametric datasets, or ANOVA with Holm-Sidak test for parametric results. All tests were performed using GraphPad Prism (GraphPad Software Inc).

As CRS is the most common side effect associated with CAR T-cell treatment, we investigated whether itacitinib is able to reduce the levels of cytokines associated with acute hyperactivation leading to CRS. Therefore, we conducted experiments in which naïve animals were challenged with ConA, a potent T-cell mitogen capable of inducing broad inflammatory cytokine releases and proliferation (26). Similar to individuals experiencing CRS, animals receiving ConA have elevated serum levels of multiple inflammatory cytokines as well as behavioral changes such as fever, malaise, hypotension, hypoxia, capillary leak, multiorgan toxicity, and potentially death. To study the effect of itacitinib in this model, corresponding animals were prophylactically dosed with 60 or 120 mg/kg of itacitinib to achieve JAK1 inhibition coverage equivalent to that observed in clinical trials (27–29). When compared with vehicle-dosed animals, itacitinib was able to significantly reduce serum levels of many of the cytokines implicated in CRS (i.e., IL6, IL12, and IFNγ) in a dose-dependent manner (Fig. 1A). As expected, itacitinib did not have a significant effect on cytokines independent of the JAK1 pathway (i.e., IL5; Fig. 1A). However, not all JAK-mediated cytokines were significantly decreased (i.e., IL4; Fig. 1A). In addition, itacitinib was also able to dose-dependently reduce CRS-implicated cytokines in a therapeutic mode, where animals were dosed with itacitinib 30 minutes after ConA challenge (Fig. 1B).

To confirm the capacity of itacitinib to reduce hyperinflammation, naïve animals were challenged with anti-CD3 to induce nonspecific T-cell activation and cytokine response. Once again, corresponding animals were either prophylactically or therapeutically dosed with 120 mg/kg of itacitinib. Compared with vehicle-treated mice, itacitinib was able to significantly reduce serum levels of many of the cytokines implicated in CRS, but have no effect on cytokines independent of the JAK1 pathway (Supplementary Fig. S1A).

Recent evidence shows that host macrophages are the major producers of IL6 after CAR T-cell treatment (30). Therefore, we queried whether itacitinib would prevent IL6 production by host macrophages. First, murine bone marrow–derived macrophages were expanded in vitro with G-CSF, and itacitinib was added to the cultures at day 6. LPS was added to the culture at day 7 to activate the macrophages (31). Prophylactic treatment with itacitinib reduced IL6 production in a dose-dependent manner, indicating that the activity of itacitinib in reducing production of inflammatory cytokines is not exclusive to T cells (Fig. 2A). We also observed a nonsignificant trend toward reduction on several other cytokines (i.e., IL10, IL12p70, KC/GRO; Supplementary Fig. S2).

Having determined that itacitinib is able to reduce IL6 production in vitro, we next expanded our studies to an in vivo context to assess the effect of itacitinib on activated macrophages in mice. Mice were prophylactically treated with itacitinib or vehicle for 3 days to achieve steady state before receiving an intraperitoneal injection of LPS. Two hours after injection, cytokines were measured from intraperitoneal lavages. As seen in Fig. 2B, doses of itacitinib that mimic the JAK1 inhibition coverage achieved in clinical trials significantly reduced IL6 production by activated macrophages in vivo. Thus, data from both in vitro and in vivo systems indicate that itacitinib is capable of downregulating the major cellular source of inflammatory IL6 during an experimentally induced model of CRS.

Whereas cytokine production by the host immune system is a primary cause of CRS, production of inflammatory cytokines by CAR T cells is also a major contributor (32). Having shown that itacitinib is capable of reducing inflammatory cytokine production by the host immune system, we next wanted to examine whether itacitinib reduces inflammatory cytokine production by CAR T cells. Human CD19-CAR T cells were expanded under concentrations of tocilizumab or itacitinib equivalent to human doses achieved in clinical trials (33). After a 3-day expansion, CAR T cells were cocultured with CD19-expressing NAMALWA target cells, and 6 hours later supernatants were collected to quantify cytokines. Itacitinib, but not tocilizumab, was able to significantly reduce the levels of many inflammatory cytokines (i.e., IL2, IFNγ, IL6, and IL8; Fig. 3). As an internal control (background), we also measured cytokines produced by nontransduced T cells (Fig. 3).

As T cells play an important antitumor role, the effect of itacitinib on T cells was studied to determine whether treatment may result in broad immunosuppression potentially interfering with normal T-cell proliferation and function. T cells were obtained from freshly isolated human PBMCs of healthy adults. Following activation with anti-CD3/anti-CD28 coated beads (23), T cells were expanded in the presence of increasing concentrations of itacitinib, and proliferation was measured by flow cytometry. When compared with the DMSO control, concentrations of itacitinib relevant to the IC₅₀ (50–100 nmol/L; ref. 34) did not have a significant effect on anti-CD3/anti-CD28–induced expansion of human T cells (Fig. 4A), demonstrating that itacitinib treatment has no negative impact on the ability of T cells to proliferate. As an internal control, and consistent with the disruption of the JAK/STAT signaling activity, higher concentrations of itacitinib (1,000 nmol/L) blocked T-cell proliferation (Fig. 4A).

To study the effect of itacitinib specifically on CAR T-cell expansion and cytolytic activity, GD2-CAR T cells (23) were expanded with anti-CD3/anti-CD28–coated beads in the presence of increasing itacitinib concentrations, ranging from 100 to 500 nmol/L. Once again, when compared with the DMSO control, itacitinib at 100–250 nmol/L did not have a significant effect on CD3/CD28-induced expansion of GD2-CAR T cells, whereas 500 nmol/L was enough to block CAR T-cell proliferation (Fig. 4B). Importantly, GD2-CAR T cells expanded in the presence of low (100 and 250 nmol/L) itacitinib concentrations were able to efficiently lyse GD2-expressing tumor cells (Fig. 4C). As a negative (background) control, we also measured target cell lysis by nontransduced (nonspecific) T cells. As expected, GD2-CAR T cells expanded in the presence of a high dose of itacitinib (500 nmol/L) and were unable to induce the lysis of target cells beyond background levels (Fig. 4C). Taken together, these data indicate that IC₅₀ relevant doses of itacitinib do not interfere with CAR T-cell proliferation or antitumor activity in vitro.

Next, we queried whether itacitinib has an effect on CAR T-cell antigen-specific proliferation. EGFR-CAR T cells (35) were expanded with anti-CD3/CD28 beads for 2 weeks and then restimulated with EGFR beads in the presence of itacitinib or DMSO controls. Once again, doses of itacitinib relevant to the IC₅₀ (100 or 250 nmol/L) did not affect CAR T-cell antigen-specific proliferation (Fig. 4D).

To further test the effect of itacitinib on antigen-specific T-cell proliferation and in vivo antitumor activity, we isolated splenocytes from OT-1 mice, transgenic for the TCR Vα2Vβ5 specific for the peptide SIINFEKL (36). OT-1 T cells were expanded with the SIINFEKL peptide in the presence of increasing concentrations of itacitinib, and their expansion was measured by flow cytometry. As expected, itacitinib concentrations relevant to the IC₅₀ have a minimal effect on the expansion rate (Supplementary Fig. S3A). Itacitinib concentrations relevant to maximum free concentrations (189 and 244 nmol/L, in the absence or presence of potent CYP3A4 inhibitors, respectively) induced a modest reduction on T-cell proliferation (Supplementary Fig. S3A). To evaluate the effect of itacitinib on the in vivo antitumor efficacy of OT-1 CD8, we conducted experiments involving adoptive transfer of OVA-specific CD8 cells into C57BL/6 mice previously transplanted with an OVA-expressing tumor cell line (EG7). Five days after tumor challenge, corresponding animals were orally dosed twice a day with vehicle or 60 or 120 mg/kg of itacitinib for 2 weeks. Eight days after tumor inoculation, animals received an adoptive transfer of OVA-specific OT-1 naïve CD8 cells. When compared with the control group, adoptive transfer of OT-1 CD8 was able to significantly reduce tumor growth (Fig. 4). Importantly, itacitinib doses equivalent to the JAK1 coverage target doses do not affect the antitumor efficacy of OT-1 cells (Fig. 5).

To rule out a direct effect of itacitinib on EG7 tumor cell growth, we expanded EG7 cells in the presence of increasing itacitinib concentration. Even very high concentrations of itacitinib (25 μmol/L) were unable to inhibit tumor proliferation (Supplementary Fig. S3B).

To study the effect of itacitinib on human CAR T cells targeting the CD19 antigen, we treated CD19-CAR T cells with either itacitinib or anti-IL6 receptor (tocilizumab) for 3 days and then measured their cytolytic activity against CD19-expressing target cells. Whereas concentrations of tocilizumab that mimic human pharmacologic activity significantly reduced the cytolytic activity of CAR T cells (37), 100 nmol/L of itacitinib showed no significant effect when compared with control CAR T cells (Fig. 6A). CAR T cells expanded with a higher dose of itacitinib (300 nmol/L, ∼5-fold higher that the cellular IC₅₀) showed a modest but statistically significant inhibition on the antitumor activity (Fig. 6A). As internal controls, the antitumor activity of nontransfected PBMCs was also tested. Nontransduced T cells were unable to specifically lyse the target cells and were unaffected by neither itacitinib nor tocilizumab treatment beyond background levels (Fig. 6B).

To test the effect of itacitinib in vivo, we studied the effect of oral itacitinib on the antitumor efficacy of CD19-CAR T cells. Immunodeficient mice (NSG) were challenged with CD19⁺ Nalm6-luciferase–expressing human lymphoma cells. Once the tumor was engrafted (day 4), the mice received an adoptive transfer of 3 × 10⁶ human CD19-CAR T cells. In this model, CAR T cells were able to control tumor growth, measured by luciferin expression (Fig. 7). Importantly, daily oral itacitinib dosing (120 mg/kg) of the animals did not affect the antitumor activity of adoptively transferred CAR T cells, thus indicating that, at targeted doses, itacitinib does not have a significant effect on the antitumor activity of CAR T cells.

Finally, to confirm that itacitinib would not affect CAR T-cell activity on a more aggressive model, immunodeficient mice (NSG) were challenged with 5 × 10⁶ CD19⁺ NAMALWA-luciferase–expressing human lymphoma cells. Once the tumor was engrafted (day 7), the mice received an adoptive transfer of human CD19-CAR T cells. When compared with animals receiving control cells, animals receiving an adoptive transfer of CAR T cells had a significant tumor growth delay, measured both by luciferin expression (Supplementary Fig. S4B) and expansion of survival (P = 0.0014; Supplementary Fig. S4C). Even in this aggressive tumor model, daily oral itacitinib dosing of the animals did not affect the antitumor activity of adoptively transferred CAR T cells (P = 0.1860), thus confirming the safety of prophylactic itacitinib dosing to prevent CRS.

CAR T-cell therapy has shown unprecedented remission rates in patients with relapsed or refractory B-cell acute lymphoblastic leukemia. Unfortunately, the high frequency of side effects (mainly CRS and neurotoxicity) that often require extensive critical care support remains a barrier to the further development of CAR T-cell therapy. Current therapeutic options are limited to treating severe cases of CRS with tocilizumab or corticosteroids, but unfortunately severe toxicities and death still occasionally occur (38). Therefore, there is a clear need to develop prophylactic treatments to prevent (rather than treat) CRS without impairing antitumor efficacy.

Despite being a very active research field, only recently clear predictors of CAR T cell–associated toxicities have been identified, and several factors, such as ferritin, C-reactive protein, or tumor burden can be reliable predictors of severe CRS (11). A recent clinical trial on pediatric patients with leukemia suggests that early therapeutic intervention with tocilizumab is effective at reducing CRS rates without affecting CAR T-cell proliferation or antitumor efficacy (39). However, these data should not be generalized to other conditions such as lymphoma as clinical management strategies are different than leukemia and concerns remain over the potential impact of corticosteroids on CAR T-cell activation and proliferation (14).

Recent advances in our understanding of the JAK/STAT pathway have highlighted its role in T-cell priming, activation, and expansion. Indeed, several small molecules targeting the JAK/STAT pathway have shown clinical potential in a number of indications (reviewed in ref. 40). Biochemical and cellular profiling of itacitinib confirmed it to be a potent JAK1 inhibitor with large fold-selectivity margins over the other family members (JAK2, JAK3, and TYK2) or unrelated kinases (41). Importantly, JAK1 either independently or in cooperation with other members of the JAK family, mediate the signaling of a number of inflammatory cytokines implicated in CRS, including GM-CSF, IFNγ, IL6, and IL12 (42, 43). Crucially, specific inhibition of JAK1 may prevent some of the side effects associated with JAK1/JAK2 inhibitors such as lymphopenia (42).

Consistent with the potent disruption of the JAK1-specific STAT signaling activity, here we show that itacitinib inhibited multiple cytokines involved in CRS (including IL6, IL12, and IFNγ) in two different models of systemic inflammation in a dose-dependent manner. Importantly, only cytokines dependent upon JAK1 were affected, as levels of IL5 and other JAK1-independent cytokines were unaffected by itacitinib treatment. It is also important to note that while the effect on inflammatory cytokine levels was profound and statistically significant, plasma levels of such cytokines were still elevated when compared with naïve animals, thus potentially dampening the hyperimmune activation state, but avoiding broad immunosuppression.

In addition to the effect of itacitinib on the production of cytokines by T cells and CAR T cells, our findings show that itacitinib is also able to reduce the secretion of inflammatory cytokines by macrophages, both in vitro and in vivo. Recent studies have shown that IL1 and IL6 produced by host monocytes/macrophages play a key role in the pathogenesis of CRS and neurotoxicity (30, 44). Our data demonstrate that itacitinib is able to inhibit IL6 production by murine macrophages in vivo, demonstrating its potential utility in treating the most prominent source of inflammatory cytokines during CRS. The ability of itacitinib to inhibit the production of CRS-associated cytokines produced by a range of cell types (both of murine and human origin, an important distinction considering the differences between mouse and human biology; ref. 45) is particularly noteworthy, as such a strategy may be necessary for the effective treatment of CRS.

To achieve clinical efficacy while preventing CRS, it is imperative that clinical doses of itacitinib do not interfere with CAR T-cell proliferation and antitumor activity. Our findings confirm that itacitinib concentrations relevant for cellular IC₅₀ (in vitro) and drug pharmacokinetic exposures that mimic JAK1 target coverage in vivo have no significant effect on the proliferation rate of PBMCs or four different CAR T-cell constructs. More importantly, our data show that in preclinical models, where animals receive an oral dose of itacitinib that mimic human JAK1 pharmacologic inhibition, prophylactic treatment with itacitinib does not affect the in vivo antitumor efficacy of CAR T cells.

In summary, we have demonstrated that itacitinib at clinically relevant doses can prevent CAR T cell–associated CRS in preclinical models without negatively affecting antitumor efficacy. Reducing the incidence and severity of CRS has the potential to benefit clinical practice, including the use of CAR T-cell therapy in patients in previously high-risk populations. On the basis of these results, a phase II clinical trial of itacitinib for the prevention of CRS induced by CD19-CAR T-cell therapy was initiated (NCT04071366). Results from our preclinical study may have implications in other settings; future studies are required to further investigate the role of JAK inhibition beyond CAR T cell–induced CRS (i.e., bispecific antibodies; ref. 46) or conditions [including HLH (9, 47) and respiratory infections such as SARS (48), MERS (49), or Covid-19 (50–52)] where CSR may play a role.

E. Huarte is an employee and stockholder of Incyte Research Institute. M.T. Peel is employed by Incyte and holds shares in Incyte. L. Truong reports personal fees from Incyte outside the submitted work. A. Naim reports other from Incyte Corp (employment, stock) outside the submitted work. M.C. Milone reports grants from Incyte during the conduct of the study and grants from Novartis outside the submitted work; in addition, M.C. Milone has a patent for CAR T-cell technology issued, licensed, and with royalties paid from Novartis. P.A. Smith reports other from Incyte Corporation (employee) during the conduct of the study. No potential conflicts of interest were disclosed bythe other authors.

E. Huarte: Conceptualization, data curation, formal analysis, methodology, writing-original draft, writing-review and editing. R.S. O'Connor: Data curation, formal analysis, methodology. M.T. Peel: Data curation, formal analysis, methodology, writing-original draft, writing-review and editing. S. Nunez-Cruz: Data curation. J. Leferovich: Data curation. A. Juvekar: Data curation, formal analysis, methodology. Y.-o. Yang: Formal analysis, methodology. L. Truong: Data curation, formal analysis. T. Huang: Formal analysis. A. Naim: Conceptualization, formal analysis. M.C. Milone: Formal analysis. P.A. Smith: Conceptualization, formal analysis, writing-original draft, writing-review and editing.

This study was funded by Incyte Corporation, and supported in part by an NIH grant RO1CA226983–03 awarded to R.S. O'Connor.

We are thankful to April Barbour, Michael Howell, and Lea Burke for useful discussions about pharmacokinetics and translational relevance of the manuscript. Editorial assistance was provided by Envision Pharma Group, Inc. (Philadelphia, PA), and funded by Incyte Corporation.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.