Programme Overview Session: FS 01

Content

Cancer vaccination using tumor antigen-primed dendritic cells (DCs) was introduced in the clinic some 25 years ago, but the overall outcome has not lived up to initial expectations. There are many factors that determine the efficacy of DC therapy. It has become clear that we need non-invasive imaging methods capable to a) verify that tumor antigen-primed DCs are administered correctly at the targeted injection site¹; b) to assure that tumor antigen-primed DCs home in sufficient numbers to the LN follicles where recipient T cells reside and are waiting to be activated; and c) that homing, tumor antigen-primed DCs indeed carry the required amount of antigen needed for sufficient T cell activation and treatment efficacy. Ideally, quantification of LN signal changes will serve as an early predictor for the efficacy of cancer vaccination and anti-tumor treatment response².

To address these uncertainties, proton (¹H) and fluorine (¹⁹F) magnetic resonance imaging (MRI) tracking of ex vivo pre-labeled DCs has been used to non-invasively determine the accuracy of therapeutic DC injection, initial DC dispersion, systemic DC distribution, and DC migration to and within LNs (Figure 1)³. Magnetovaccination is an alternative approach that tracks in vivo labeled DCs that simultaneously capture tumor antigen and MR contrast agent in situ, enabling an accurate quantification of antigen presentation in LNs (Figure 2)⁴. The latter approach mimics the natural biological process of DC-tumor cell interactions, is more specific than labeling DCs ex vivo, and can visualize the subset of true antigen-presenting DCs. The magnetovaccination strategy has been used to study the timing and dosing of immunoadjuvants such as the Toll-like receptor 4 (TLR4) agonist glucopyranosyl lipid A (GLA). It was shown that injection of GLA 24 hours prior or simultaneously with the injection of DCs paradoxically decreased the amount of SPIO-labeled DCs homing to LNs, but this could be rescued by injecting GLA 24 hours after the injection of DCs⁵. It was postulated that the immunoadjuvant has such a potency that it induces DC LN homing before these cells have sufficient time to capture the tumor antigen and SPIO in situ.

Recently, instead of DCs priming T cells in vivo, autologous cytotoxic T cells have been isolated and engineered in vitro to express a receptor against a tumor antigen. These so-called chimeric-antigen receptor (CAR T cells) were approved by the FDA for acute lymphoblastic leukemia in the summer of 2017, “ushering in a new approach to the treatment of cancer” at a staggering cost of nearly $500,000 for a one-time treatment. Since then, other types of cancer have been targeted, some with remarkable treatment success, while others did not respond. A concern is that life-threatening side effects have occasionally been reported, i.e., cytokine release syndrome, a systemic response not unlike the ravaging cytokine storm our body can unleash following contraction of COVID-19. Clearly, non-invasive imaging approaches that can report on the efficacy of CAR T cell therapy early on are warranted, including their tumor homing⁶.

As a concluding remark it should be mentioned that the number of clinical MRI immune cell tracking studies have been few⁷, which may have its roots in the paucity of available GMP-grade labeling agents, their cost of production and testing, and the need for cumbersome IND approval for their off-label use (unlike the use of radionuclides, where often the microdosing tracer principle applies). Chemical exchange saturation transfer (CEST) MRI tracking of unlabeled cells would avoid these hurdles, but has so far only be applied for mesenchymal stem cells⁸.

References

1.         de Vries, I. J.M. et al., Nat Biotechnol 2005, 23 (11), 1407-13.

2.         Bulte, J. W. M. et al.,  Mol Imaging Biol 2022 in press (DOI 0.1007/s11307-021-01647-4).

3.         Ahrens, E. T. et al,  Nat Rev Immunol 2013, 13 (10), 755-63.

4.         Long, C. M. et al., Cancer Res 2009, 69 (7), 3180-7.

5.         Kadayakkara, D. K. et al. Cancer Res 2015, 75 (1), 51-61.

6.         Kiru, L. et al., Proc Natl Acad Sci U S A 2022, 119 (6).

7.         Bulte, J. W. M. et al., Radiology 2018, 289 (3), 604-615.

8.         Yuan, Y. et al. Nat Biomed Eng 2022 in press (doi 10.1038/s41551-021-00822-w).

Disclosure

a) I or one of my co-authors have no financial interest or relationship to disclose regarding the subject matter of this presentation.

Content

Fluorescence molecular imaging (FMI) is an innovative imaging modality that improves the way in which we detect a wide variety of diseases and potentially guide treatment follow-up. Innovation in detection can be conceived in the broader sense of the word; from detection of white light invisible malignant lesions or inflammation, to elucidating drug distribution enabling prediction of therapy response and determining optimal dose concentration in individual patients.

In FMI, a near-infrared fluorescent dye is coupled to a substrate, such as an antibody or peptide, to selectively bind to proteins that are overexpressed in the tissue or elucidate drug distribution. In addition to the qualitative visualization of the fluorescence, we can also accurately quantify the fluorescent intensity using spectroscopy. The in vivo fluorescent measurements are correlated to ex vivo fluorescent imaging, immunohistochemistry and fluorescence microscopy giving detailed insight in for instance local drug distribution and binding of target cells.

We have performed multiple clinical trials in oncology and inflammatory bowel disease (IBD) to reveal the multiple purposes for which FMI can be beneficial during endoscopy. These range from improved detection of early lesions in esophageal carcinoma, to detection of residual disease in rectal cancer after neoadjuvant treatment, to elucidating drug distribution in IBD patients. Within IMI we aim to improve insight in anti-PD-L1 drug distribution before and during neoadjuvant therapy in esophageal cancer.  In our presentation examples and challenges of FMI will be highlighted.

Acknowledgement

Disclosure

I or one of my co-authors have no financial interest or relationship to disclose regarding the subject matter of this presentation.

Content

Introduction

CD8-positive T cells play an important role in the antitumor response. Monitoring CD8 T cells could provide valuable patient selection and treatment outcome information. To preclinically monitor CD8 T cells, we developed a novel nanobody specifically targeting murine CD8 and radiolabeled this nanobody with fluorine-18 to allow same-day PET imaging.

Materials and methods

Llama immunization with murine CD8 followed by phage display, resulted in several anti-CD8 nanobody clones. We characterized the CD8 nanobody in vitro and determined affinity by surface plasma resonance to select a high-affinity candidate. The selected nanobody included a LPETG motif to allow site-specific conjugation using sortase. We site-specifically conjugated the nanobody with tetrazine to allow ¹⁸F-transcyclooctene radiolabeling. We then further characterized the nanobody and measured immunoreactive fraction by size-exclusion HPLC and impact on T cell survival and proliferation by flow cytometry. Next, we studied ¹⁸F-CD8 nanobody in vivo in nontumor- and tumor-bearing mice by PET imaging up to 2 hours after intravenous tracer administration. In nontumor-bearing mice we assessed dose-dependent uptake and specificity by including a nonbinding control nanobody. Two tumor-bearing mice models were used. First a HER2+ Founder5 (Fo5) xenograft model was used to study T cells dynamics upon a T cell-dependent bispecific (TDB) HER2/CD3 antibody treatment. Second, CT26 xenograft-bearing mice were treated with a combination of an anti-PD-L1 checkpoint inhibitor and a MEK inhibitor (MEKi). Both tumor-bearing mouse models were scanned prior to treatment and post-treatment to assess CD8 T cell dynamics. Aspecific tumor uptake was assessed by including an amino acid matched nonbinding control nanobody.

Results

We successfully conjugated and radiolabeled the CD8 nanobody. In vitro, CD8 nanobody showed subnanomolar affinity to murine CD8, an immunoreactive fraction >90% and did not impact T cell proliferation and survival. In nontumor-bearing mice, ¹⁸F-CD8 nanobody specifically visualized primary and secondary lymphoid tissues, including spleen, thymus and lymph nodes. No uptake was observed in these tissues with ¹⁸F-labeled control molecule. Increasing tracer dose of ¹⁸F-CD8 nanobody resulted in a dose dependent decrease in lymph nodes and spleen uptake. Imaging the mice of the highest dose group the next day visualized spleen and lymph nodes, indicating that repeat imaging within 24 hours was possible. In Fo5 xenograft-bearing mice, pre-treatment ¹⁸F-CD8 nanobody levels in the tumor were low. However, 4 days after a single dose of HER2/CD3 TDB, PET imaging revealed a significant increase in ¹⁸F-CD8 nanobody levels.  Flow cytometry post treatment confirmed the CD8 T cell infiltrate in these Fo5 tumors. ¹⁸F-labeled nonbinding control nanobody showed no difference in tumor uptake between pre- and post-TDB. In the second tumor model, baseline levels of ¹⁸F-CD8 nanobody in CT26 xenografts were low. Seven days post MEKi + anti-PD-L1 treatment, tumor levels of ¹⁸F-CD8 nanobody were increased significantly whereas the nonbinding control molecule showed no difference in tumor uptake upon combination treatment.

Conclusion

CD8-nanobody has shown to detect CD8 T cells in physiological tissues as well as changes in CD8 tumor levels upon cancer immunotherapy treatment. CD8 nanobody imaging has the potential to be used as a biomarker to assess treatment response.

Acknowledgement

Disclosure

I have the following financial interest to disclose the subject matter of this presentation: Employment at Genentech Inc.