Cancer recurrence after surgical resection remains a significant challenge in cancer therapy. Platelets, which accumulate in wound sites and interact with circulating tumour cells (CTCs), can however trigger inflammation and repair processes in the remaining tumour microenvironment. Inspired by this intrinsic ability of platelets and the clinical success of immune checkpoint inhibitors, here we show that conjugating anti-PDL1 (engineered monoclonal antibodies against programmed-death ligand 1) to the surface of platelets can reduce post-surgical tumour recurrence and metastasis. Using mice bearing partially removed primary melanomas (B16-F10) or triple-negative breast carcinomas (4T1), we found that anti-PDL1 was effectively released on platelet activation by platelet-derived microparticles, and that the administration of platelet-bound anti-PDL1 significantly prolonged overall mouse survival after surgery by reducing the risk of cancer regrowth and metastatic spread. Our findings suggest that engineered platelets can facilitate the delivery of the immunotherapeutic anti-PDL1 to the surgical bed and target CTCs in the bloodstream, thereby potentially improving the objective response rate.
Surgery is the main treatment option for most solid tumours. Despite continual improvements in surgical techniques, residual microtumours and/or circulating tumour cells (CTCs) after tumour resection remain a challenge
One mechanism that may account for the compromised efficacy of anti-PD therapy is off-target binding of antibodies to normal tissues when intravenously infused
. In this context, it is ideal for cancer immunotherapy to focus on the diseased site whilst preventing the systemic nonspecific over-activation of the immune system
. The limited efficacy may also be due to insufficient lymphocytic infiltration of the tumour and insufficient T-cell-induced inflammation in the tumour microenvironment
Cell-based systems have recently emerged as biological drug carriers; examples include erythrocytes, bacterial ghosts, and genetically engineered stem and immune cells
. Among them, platelets are anucleated cellular fragments released from megakaryocytes and are best known for their function in hemostasis
In this work, inspired by the intrinsic properties of platelets, we conjugated anti-PDL1 (antibodies against PDL1; hereafter, aPDL1) to the surface of platelets for use as a preventative treatment for post-surgical tumour recurrence (Fig. 1a). With the help of platelets, aPDL1 can be targeted to the cancer cells after surgery, while reducing off-target effects. We found that the binding of aPDL1 to non-activated platelets was highly stable, while release of aPDL1 could be significantly promoted on the activation of platelets. We reasoned that the aPDL1 release may result from the platelet-derived microparticles (PMPs), which are generated from the plasma membrane of activated platelets 43 . Such structural alterations can facilitate aPDL1 binding to tumour cells and APCs. By intravenous injection of aPDL1-conjugated platelets (P–aPDL1) into mice with B16 melanomas and triple-negative mammary carcinomas that had been previously resected (~1% remaining), we showed that platelets can facilitate aPDL1 transport to residual microtumours at the surgical site, and to CTCs in the blood. A T-cell-inflamed tumour microenvironment was created by the platelets on activation, leading to increased PDL1 expression at the tumour site. aPDL1 was effectively released after platelet activation, thereby blocking PDL1 on tumour and APCs. Our results show that platelets have promise as a means of targeted, controlled delivery of aPDL1 for the prevention of cancer recurrence post surgery.
Engineering platelets decorated with aPDL1
A covalent approach was employed to conveniently conjugate aPDL1 to platelets via a bifunctional maleimide linker, which bypasses the need for genetic modification 44 . The binding of aPDL1 to platelets was validated using immunofluorescence (Supplementary Fig. 1). In addition, the maleimide-free aPDL1 showed minimal nonspecific binding to the platelets (Supplementary Fig. 2). The enzyme-linked immunosorbent assay (ELISA) showed that platelets could be readily coupled with aPDL1, at up to ~0.3 pg per platelet (Supplementary Fig. 3a). We found that coupling 0.1 and 0.2 pg of aPDL1 per platelet had no obvious effect on their viability (Supplementary Fig. 3b). Such covalent binding was highly stable, and did not show significant release within two days (Supplementary Fig. 3c). Furthermore, several platelet surface proteins, which modulate cell adhesion and migration, were examined after platelet decoration and we did not find any significant changes in these proteins (Supplementary Fig. 4). Overall, conjugation of aPDL1 did not induce any significant damage in the decorated platelet cells.
Activation of platelets promotes aPDL1 release
Platelet activation happens after adhesion occurs 45 . A platelet contains about 60 granules, which in turn contain many molecules with immune functions 38 . On platelet activation, the granules release these immunoactive molecules into the extracellular environment, leading to recruitment and activation of other immune cells, and induction of T-cell migration and monocyte differentiation into dendritic cells. In addition, PMPs are generated from the plasma membrane on platelet activation 43 : activated platelets release PMPs carrying adhesion molecules and chemokines, facilitating monocyte traps at the site of PMP deposition 46 . To examine whether P–aPDL1 could be activated on stimulation, we used thrombin to activate them in vitro. Transmission electron microscopy (TEM) images of P–aPDL1, before and after activation, indicated that large numbers of PMPs were generated (Fig. 1b, Supplementary Fig. 5). It was also observed that the platelet morphology became more dendritic and expansive. More interestingly, significant release of aPDL1 from the activated platelets was found using ELISA (Fig. 1c). The remarkable aPDL1 release was ascribed to PMPs that were dissociated from the plasma membrane on platelet activation. To test this hypothesis, immunofluorescence imaging of activated P–aPDL1 was carried out. Platelets were stained with calcein and aPDL1 was stained with a fluorescent secondary antibody (Supplementary Fig. 6). It was found that the aPDL1 was present on the PMPs after activation of platelets.
Several pro-inflammatory cytokines were released with aPDL1 after platelet activation (Fig. 1d,e). To further validate the aPDL1 release from the platelets via PMP generation, a transwell culture system was used. P–aPDL1 were cultured in the upper compartment of the transwell culture system, while B16 cancer cells were grown in the lower compartment (Supplementary Fig. 7). The micropores in the insert membrane had a diameter of 1 μm, allowing PMPs to cross the membrane freely. The cancer cells were co-incubated with non-activated and activated platelets for 12 h before immunostaining. As shown in Fig. 1f and Supplementary Fig. 7, a large amount of aPDL1 was bound to the cancer cell membranes where the platelets had been activated; whereas, very little binding (fluorescence) was detected in the wells with non-activated platelets. Collectively, the data suggest that aPDL1 is released from activated platelets and that it binds to cancer cells.
In vivo P–aPDL1 therapy
Coupling of aPDL1 to the surface of platelets has the potential to alter their in vivo behaviour. Therefore, the in vivo pharmacokinetics of P–aPDL1, following systemic administration, was evaluated in healthy mice. A solution of free aPDL1 or a suspension of P–aPDL1, or a mixture of unconjugated platelets and free a-PDL1, were intravenously injected into the mice using equivalent aPDL1 doses. ELISA analysis of the blood showed that the circulation half-life for aPDL1 was greatly increased when it was linked to the platelets: 34.8 h versus 5.2 h and 5.5 h for the free-aPDL1 solution and the mixture, respectively (Fig. 2a). The relatively short blood-circulation time compared with other therapeutic antibodies in clinical use is attributable to the higher immunogenicity and nonspecific binding of the rat anti-mouse IgG. We then tested their capacity to accumulate at the wound site, after incomplete removal of the primary tumour by surgery. aPDL1 was fluorescently labelled using Cy5.5 and then conjugated to platelets. In vivo fluorescence imaging was conducted 2 h post injection of the P–aPDL1 or free aPDL1, with major organs collected for ex vivo fluorescence imaging (Supplementary Fig. 8). It was observed that aPDL1 was enriched around the surgical wound and residual tumours when it was conjugated to the platelets, whereas insignificant fluorescence signal was detected at the wound site for the free aPDL1 (Fig. 2b). Ex vivo imaging of major organs, as well as the wounds with residual microtumours, further confirmed the accumulation of P–aPDL1 at the target sites (Fig. 2c, Supplementary Fig. 8). The predominant aPDL1 fluorescence was in the liver, but the P–aPDL1 showed significantly lower fluorescence here despite the equivalent dose, suggesting a longer blood circulation time for P–aPDL1. Moreover, P–aPDL1-treated mice showed 9.4-fold greater fluorescence around the surgical wound and residual microtumours than those treated with the free aPDL1; at 2 h post intravenous (i.v.) injection (Fig. 2d). Confocal images of microtumour slices also indicated significantly increased tumour uptake of aPDL1 when conjugated to platelets (Fig. 2e, Supplementary Fig. 9a). Furthermore, aPDL1 released from platelets was clearly observed in vivo via fluorescence imaging (Supplementary Fig. 9b). The haemostatic effect of modified platelets was also examined, and we found that mouse tail bleeding time did not significantly differ between mice receiving P–aPDL1 and those receiving undecorated (naïve) platelets (Supplementary Fig 10).
To examine the effect of platelet injection on local and systemic inflammation in vivo, we assessed the levels of pro-inflammatory cytokines after surgical removal of the primary tumour, and i.v. injection of platelets. Six hours post injection, wound tissue and serum were collected, and the tissue was cultured in medium for 24 h. After examination of this medium, it was found that levels of the cytokines, IL1β, IL6, TNFα and sCD40L, were all elevated versus those of the untreated-control (no surgery) and wound-bearing (surgery alone) mice (Supplementary Fig. 11a). In contrast, the serum levels were not increased by the platelets (Supplementary Fig. 11b), so our results suggest that the injection of platelets induced a local pro-inflammatory environment at the surgical site, without causing systemic inflammation. A pro-inflammatory tumour environment facilitates aPDL1 immunotherapy by converting quiescent precursor lymphocytes into activated lymphocytes 47 . Consistent with this, we found that expression of PDL1 was upregulated at the tumour site in both the tumour cells and tumour-infiltrating immune cells, perhaps as a result of the platelet-induced pro-inflammatory environment (Supplementary Fig. 11c,d). The increased number of PDL1-positive cells within the tumour should further enhance the anti-PDL1 immunotherapy and potentially increase the objective response rate 41,42 .
To investigate P–aPDL1 treatment of residual microtumours that remain after surgery, we used the B16F10 mouse melanoma incomplete-tumour-resection model to mimic post-surgical local recurrence (Fig. 2f). After surgically removing most of the tumour (~99%), the mice were intravenously injected with a single dose of phosphate-buffered saline (PBS), platelets, aPDL1 or P–aPDL1 (aPDL1 = 1 mg kg−1). Tumour growth was monitored via the bioluminescence of B16F10 cells. We found that the mice receiving P–aPDL1 exhibited the smallest relapsed-tumour volumes, with 6 of 8 in this group showing strong responses without any detectable tumour. In contrast, those treated with free aPD1 showed a modest delay of tumour growth but recurrence was not prevented in the surgical bed. When unconjugated platelets alone were used, tumour recurrence was similar to that in the PBS controls (Fig. 2g,h). The mouse survival rate, which was correlated with tumour size (Fig. 2i), was about 75% after 60 days for mice receiving P–aPDL1. In stark contrast, no mice survived beyond day 30 for all other groups (Fig. 2j).
T-cell-mediated immune responses
Tumour-infiltrating lymphocytes (TILs) from the recurrence tumours were harvested and analysed via immunofluorescence and flow cytometry on day 16. Immunofluorescence staining revealed that the residual tumours in the control group had limited T-cell infiltration. In contrast, those from P–aPDL1-treated mice were highly infiltrated by both CD8+ and CD4+ T cells (Fig. 3a, Supplementary Fig. 12). The tumour weights were significantly lower in the P–aPDL1-treated mice on day 16 (Fig. 3b), which was associated with an increase in the absolute number of CD3+ cells in the residual tumour (Fig. 3c). More importantly, the absolute number of CD8+ T cells, per gram of tumour, increased by almost tenfold in the P–aPDL1-treated mice versus the PBS controls, and threefold over the free-aPDL1-treated mice (Fig. 3d–f).
Tumour-infiltrating CD4+ Foxp3+ T cells were also examined (Fig. 3e). Intratumoural ratios of effector to regulatory T cells were significantly enhanced in mice after P–aPDL1 therapy (Fig. 3f–h). In addition, proliferation of CD8+ and CD4+ effector T cells was high within the tumours of P–aPDL1-treated mice, as measured by the expression of the cell-cycle associated protein, Ki67. In contrast, P–aPDL1 therapy did not significantly increase proliferation of tumour-infiltrating regulatory T cells (Supplementary Fig. 13). Taken together, these observations suggested that P–aPDL1 could effectively deliver aPDL1 to the tumour microenvironment, triggering a robust T-cell-mediated anti-tumour immune response.
P–aPDL1 therapy for metastatic disease
To examine the potential of P–aPDL1 to treat CTCs, we first tested it in an experimental metastasis model by intravenously injecting mice with B16F10 cells after surgery, thereby mimicking the escape of CTCs from the primary tumour and into circulation 48 (Fig. 4a, Supplementary Fig. 14). The mice were injected (i.v.) with a single dose of PBS, platelets, aPDL1 (aPDL1 = 1 mg kg−1 or 2 mg kg−1) or P–aPDL1 (aPDL1 = 1 mg kg−1), immediately after surgery. Based on the bioluminescence of the B16F10 cells in the mice (Fig. 4b, Supplementary Fig. 14), it was found that the free-aPDL1 treatment reduced metastatic cancer, but did not significantly reduce local tumour recurrence at the surgical site, even at the higher dose of 2 mg kg−1; this was possibly due to the poor accumulation of aPDL1 in and around these residual microtumours after surgery (Supplementary Fig. 8). In contrast, P–aPDL1 therapy substantially reduced both local tumour recurrence and lung metastasis; as was confirmed by observation of the whole lungs (Fig. 4c), hematoxylin and eosin (H&E) staining (Fig. 4d) and fluorescence imaging (Supplementary Fig. 15). The average number of metastatic sites in the lungs decreased greatly when the P–aPDL1 treatment was given (Fig. 4e). Furthermore, the survival time of the treated mice was significantly increased when compared with the control group (Fig. 4f). The improved anticancer effects of P–aPDL1 versus free aPDL1 can be partly attributed to the increased local concentration of antibodies around the cancer cells (Supplementary Fig. 8). Another component of the enhanced effect was platelet activation, as it not only induced release of the conjugated aPDL1, but also helped recruit many other immune cells to infiltrate the tumour microenvironment. With the PDL1 blockade, these immune cells could induce strong anticancer immune responses.
To further verify whether in situ activation of P–aPDL1 contributed to the anticancer effects, PMPs were collected from the platelets and modified with aPDL1 (Supplementary Figure 14.). Direct injection of these microparticles (separate from the platelets) clearly limited the anticancer effect, so that it was no greater than that for free-aPDL1 (Fig. 4). This could be explained by previous studies reporting that PMPs can be cleared rapidly following introduction into the circulation 49,50 . These results suggest that in situ activation of P–aPDL1 at the tumour site was an essential component of its anticancer effect.
P–aPDL1 therapy for recurrence of triple-negative carcinoma
To assess the effectiveness of P–aPDL1 in reducing another type of cancer recurrence after surgery, we performed tests in a triple-negative breast cancer (TNBC) tumour model (4T1 carcinoma). BABL/c mice were subcutaneously injected with triple-negative 4T1 tumour cells, and 14 days later, the primary tumour was surgically removed, leaving residual microtumours behind (~1% of original tumour mass). Therapeutic platelets decorated with aPDL1 were intravenously injected into the mice immediately after surgery. In this model, a single course of P–aPDL1 therapy was sufficient to reduce the growth of the residual tumour (Fig. 5a,b). Furthermore, the therapy was also very effective on lung metastases: only a few metastatic foci (median, ~2) were found in the lungs of the P–aPDL1 treated mice, whereas a median of 16 foci were found in the free-aPDL1 group, and the medians for the platelet-treated and PBS groups were ~30 (Fig. 5c,d). Mice receiving P–aPDL1 therapy after surgery also benefitted greatly in terms of their survival rate, with 75% of them still alive 60 days post tumour inoculation (Fig. 5e). In contrast, the PBS and platelet groups hit 0% around day 40, while the aPDL1 mice lasted longer but were all dead by day 60.
In summary, our strategy used in situ activation of platelets to enhance the delivery and effectiveness of aPDL1 in mice, thereby greatly reducing the presence of residual tumour cells after surgery, and hence also disease recurrence. Such a P–aPDL1 therapy could substantially improve the effectiveness of surgical interventions by reducing the risk of cancer recurrence and metastasis after resection of the primary tumour. Beyond the specific use of platelets for aPDL1 delivery, this method may inspire new treatments that apply bioparticulates for targeted delivery and bioresponsive release of therapeutics 51 .
The experiments did not use a method of randomization. The investigators were not blinded to allocation during experiments and outcome assessment.
The mouse melanoma cell line, B16F10, and mouse mammary carcinoma cell line, 4T1, were purchased from the American Type Culture Collection. B16F10-luc-GFP and 4T1-luc-GFP cells were gifts from L. Huang at the University of North Carolina at Chapel Hill. The B16F10 cells were maintained in Dulbecco’s modified Eagle medium (Gibco, Invitrogen) supplemented with 10% fetal bovine serum (Invitrogen), 100 U ml−1 penicillin (Invitrogen), and 100 U ml−1 streptomycin (Invitrogen). The 4T1 cells were maintained in RPMI-1640 medium (Gibco, Invitrogen) supplemented with 10% fetal bovine serum (Invitrogen), 100 U ml−1 penicillin (Invitrogen) and 100 U ml−1 streptomycin (Invitrogen). Master and working cell banks were generated immediately on receipt. The third and fourth passages were used for tumour experiments. Cells were tested every three months for potential mycoplasma. Re-authentication of cells was not performed after receipt.
C57BL/6 mice and BALB/c mice were purchased from Jackson Lab. Age-matched (6–10 weeks) female animals were used throughout all experiments. We performed all mouse studies in the context of the animal protocol approved by the Institutional Animal Care and Use Committee at the University of North Carolina at Chapel Hill and North Carolina State University. Experimental group sizes were approved by the regulatory authorities for animal welfare after being defined to balance statistical power, feasibility and ethical aspects. All mice were kept in accordance with federal and state policies on animal research at the University of North Carolina at Chapel Hill and North Carolina State University.
The anti-PDL1 antibody (aPDL1) used in vivo was purchased from Biolegend (cat. no. 124329, Clone: 10F.9G2). The primary antibodies used for immunostaining were against CD3 (Thermo Fisher Scientific, cat. no. A18644), CD4 (Thermo Fisher Scientific, cat. no. A18667), CD8 (Thermo Fisher Scientific, cat. no. A18609), PD1 (Biolegend, cat. no. 135227), CD11c (Biolegend, cat. no. 117309), PDL1 (Biolegend, cat. no. 124311), CD20 (Biolegend, cat. no. 150411), CD11b (Biolegend, cat. no. 101211), CD9 (Biolegend, cat. no. 124805), CD41 (Biolegend, cat. no. 133905), CD61 (Biolegend, cat. no. 104307), CD62P (Biolegend, cat. no. 148305), CD40L (Biolegend, cat. no. 106505), intracellular Ki67 (Biolegend, cat. no. 652405), and intracellular Foxp3 (eBioscience, cat. no. 71-5775-40) for fluorescence-activated cell sorting (FACS analysis following the manufacturers’ instructions. The stained cells were analysed on a CytoFLEX flow cytometer (Beckman), using the FlowJo software package (version 10.0.7; TreeStar, USA, 2014). The secondary antibodies used for immunostaining were goat anti-rat IgG (H + L; Thermo Fisher Scientific, cat. no. A18866), rabbit anti-rat IgG (H + L; Thermo Fisher Scientific, cat. no. A18920) and goat anti-rat IgG (minimal cross-reactivity; Biolegend, cat. no. 405408).
Preparation of aPDL1 conjugated platelets
Murine platelets were isolated as described previously 52 . In brief, whole blood was collected from the C57BL/6 or BALB/c mice (non-terminal collection from the orbital sinus or saphenous vein; 20 mice were used) into a plastic syringe containing 1.0 ml citrate-phosphate-dextrose (16 mM citric acid, 90 mM sodium citrate, 16 mM NaH2PO4, 142 mM dextrose, pH 7.4) and centrifuged at 100 g for 20 min at room temperature. The platelet-rich plasma (PRP) was transferred to a separate tube using a transfer pipette (wide orifice), and PGE1 was added to each tube at a final concentration of 1 μM. (Note that if the PRP has a reddish colour, discard these samples.) Platelets were isolated from the PRP via centrifugation at 800 g for 10 min. The plasma was discarded, and the platelets were resuspended carefully and slowly in Tyrode’s buffer (134 mM NaCl, 12 mM NaHCO3, 2.9 mM KCl, 0.34 mM Na2HPO4, 1 mM MgCl2, 10 mM HEPES, pH 7.4) or PBS with PGE1 added at 1 μM. Note that this buffer was released slowly along the tube wall, while minimizing agitation. Each in vivo injection required 500–600 μl whole blood.
The surface of the platelets was functionalized with aPDL1 in three steps. First, 100 μl of platelets (1 × 108) was resuspended in 400 μl of PBS (pH 8), including PGE1 (1 μM), and incubated with 0.1 mg ml−1 Traut’s Reagent (2-iminothiolane; Pierce) for 30 min at room temperature. After 30 min of reaction, the excess Traut’s Reagent was removed by centrifugation at 800 g for 10 min and washed with Tyrode’s buffer (with PGE1 added at 1 μM) three times (without resuspension to avoid unnecessary platelet activation). In the meantime, aPDL1 was mixed with sulfosuccinimidyl-4-(N-maleimidomethyl)- cyclohexane-1-carboxylate (Sulfo-SMCC, Pierce) in PBS (pH 7.4) at a molar ratio of 1:1.2 for 2 h at 4 °C. The excess Sulfo-SMCC was removed using a centrifugal filter device (molecular weight cut-off = 10 kDa) to purify the SMCC activated antibody. Lastly, platelets and antibodies were mixed in Tyrode’s buffer (+ PGE1 at 1 μM). After 2 h at room temperature, the excess antibodies were removed by centrifugation at 800 g for 10 min. The precipitate fraction was retained and washed with Tyrode’s buffer (+ PGE1 at 1 μM) twice. Platelet recovery was higher than 80% after conjugation based on the platelet count analysis. The obtained aPDL1–platelets were stored in Tyrode’s buffer (+ PGE1 at 1 μM) at room temperature prior to use in experiments. Unconjugated platelets were not separated from conjugated platelets. The amount of aPDL1 conjugated to the platelets was measured via ELISA (rat IgG total ELISA kit, eBioscience, cat. no. 88-50490-22). Freshly isolated platelets were used within 6 h. The platelet-activation marker, CD62P, was used for evaluating platelet activation. All platelet manipulations were performed at room temperature.
To study the conjugation efficiency, various amounts of aPDL1 were added to the platelets for conjugation. Unconjugated aPDL1 was removed with the supernatant after centrifugation at 800 g for 10 min. The P–aPDL1 pellet was then washed twice with Tyrode’s buffer (+ PGE1 at 1 μM), using centrifugation at 800 g for 10 min. After that, the P–aPDL1 was resuspended in 100 μl deionized water and ultrasonicated to lyse the platelets and release the aPDL1. The amount of aPDL1 conjugated to the platelets was measured using ELISA. The efficiency of aPDL1 conjugation to platelets (conjugated aPDL1 as a percentage of total aPDL1 added) was about 75% when 0.2 pg aPDL1 per platelet was added. To assess the stability of the platelets after decoration with aPDL1, platelet counts were determined at 0 h and 24 h. To study the stability of aPDL1 on platelets over time, the platelets were stored in Tyrode’s buffer (+ PGE1 at 1 μM) at room temperature. At different times, 50 μl P–aPDL1 was extracted. Released aPDL1 in the supernatant was removed by centrifugation at 800 g for 10 min. The conjugated platelets were suspended in 100 μl deionized water and ultrasonicated to release the aPDL1. The amount of aPDL1 conjugated to the platelets was measured via ELISA. To activate the platelets, 0.5 U thrombin ml−1 were added to the platelet suspension. PGE1 was removed prior to platelet activation. Platelets were activated for 30 min at 37 °C.
PMPs were prepared from platelets as previously described 53 . Before the experiments, platelet concentrates were activated by thrombin (2 U ml−1) for 30 min and centrifuged at 800 g for 10 min. The PMP-enriched supernatants were then collected and examined via antibodies for mouse aIIbb3 and CD62P (P-selectin) combined with flow cytometry. Next, aPDL1 was conjugated to the PMPs as described above. The conjugation efficiency and stability were examined using ELISA.
Transmission electron microscopy
To prevent morphological changes occurring in the subsequent preparation steps, the platelets were fixed by suspending them in 10% buffered formaldehyde solution. Immediately afterwards, the samples were centrifuged at 800 g for 10 min at room temperature. After discarding the supernatant, platelets were fixed with 2.5% glutaraldehyde in the cacodylate buffer (pH 7.2) for 90 min at 4 °C (for activated platelets, the supernatant were also collected for TEM imaging). After fixation, the platelets were washed twice using centrifugation at 800 g for 10 min at 4 °C. They were then sequentially stained with 2% uranyl acetate and lead citrate for 5 min each and then transferred to a copper grid 54 . The TEM images were obtained using a JEOL 2000FX instrument at 80 kV.
aPDL1 and cytokine release from platelets
To activate P–aPDL1, 0.5 U thrombin ml−1 was added to the suspension (~1 × 108 platelets in 500 μl Tyrode’s buffer; n = 3) for 30 min at 37 °C. At different times, 50 μl samples of P–aPDL1 were extracted. Supernatants containing aPDL1 and cytokines released from the platelets were collected by centrifugation at 800 g for 10 min. Non-activated P–aPDL1 was used as a control. The amount of aPDL1 and cytokines in the supernatants was measured using ELISA (eBioscience, cat. Nos. 88-50490-22 (rat IgG), 88-7013-22 (IL1β), 88-7064-22 (IL6), 88-7324-22 (TNF-α), BMS6010 (sCD40L)). The absorbance was read on an Infinite 200 PRO plate reader.
In vivo pharmacokinetics
Three mice were intravenously injected with free aPDL1, P–aPDL1 or unconjugated platelets + a-PDL1 mixture (aPDL1, 2 mg kg−1, platelets, 2 × 108, in 200 μl PBS for each mouse). A 10 μl sample of blood was extracted from the tail at different time points using an anticoagulation tube. Each sample was dissolved in 100 μl water (Sigma, cat. no. W4502) and treated with ultrasound for cell lysis to release the conjugated aPDL1. The aPDL1 was measured by rat IgG total ELISA kit (eBioscience, cat. no. 88-50490-22). For the in vivo biodistribution study, after removal of primary tumour with ~1% residual tissue left behind, the mice were intravenously injected with Cy5.5-labelled free aPDL1 or P–aPDL1. In vivo fluorescence images were recorded using an IVIS Spectrum system (Perkin Elmer; manufacturer-excitation/emission filters for Cy5.5; exposure time = 1 s). For ex vivo imaging, treated mice were sacrificed at 2 h post injection. Major organs and tissues were collected and imaged using the same IVIS system and settings.
In vivo tumour models
To measure the effects on cancer recurrence, 7 days after 1 × 106 B16F10 (or 4T1) or luciferase-tagged B16F10 (or 4T1) tumour cells were transplanted into the right flank of mice; the resulting tumours were ~300 mm3 in volume. These tumours were then resected, leaving about 1% residual tumour tissue behind to mimic the presence of residual microtumours in the surgical bed 3 . Briefly, animals were anesthetized in an induction chamber using isoflurane (up to 5% for induction; 1–3% for maintenance), and anaesthesia was maintained via a nose cone. The tumour area was clipped and aseptically prepped. Sterile instruments were used to remove approximately 99% of the tumour. The wound was closed using an Autoclip wound closing system. For the experimental metastasis model, 1 × 105 luciferase-tagged B16F10 (or 4T1) tumour cells in 200 μL PBS were intravenously infused into mice via the tail vein after resection of the primary tumour. The mice were weighed and randomly divided into four groups of eight mice (n = 8). Immediately after surgery, the mice were intravenously injected with one of the treatment formulations: PBS, platelets (1–2 × 108), aPDL1 (1 mg kg–1) and P–aPDL1 (1 mg kg–1 aPDL1 on 1–2 × 108 platelets); the platelets were freshly prepared from healthy mice of the same strain. The tumour burden was monitored via the bioluminescence of the cancer cells. The mice were clipped and shaved using a depilatory cream before imaging. Images were taken using an IVIS Lumina imaging system (Perkin Elmer). Tumour size was measured with a digital calliper. The tumour volume (mm3) was calculated as (long diameter × short diameter2)/2. Metastatic burden was assessed on the basis of the bioluminescence. The lungs were weighed and micrometastases were counted. Animals were euthanized when exhibiting signs of impaired health or when the volume of the tumour exceeded 2 cm3.
For the metastatic lung tumour in Fig. 5c, India ink was used to better visualize lung metastases following a standard protocol 55 . The mice were sacrificed and tumour burden was quantified (unblinded) after intratracheal injection of the ink (85 ml H2O, 15 ml ink, two drops of ammonia water) and fixation with Fekete’s solution (5 ml 70% ethanol, 0.5 ml formalin, 0.25 ml glacial acetic acid). After 2–6 h, tumour lesions were bleached whereas normal lung tissue remained stained.
In addition, the tumours were dissected from the mice after treatment and snap frozen in optimal cutting temperature (OCT) medium for immunofluorescence staining. Fluorescently labelled secondary antibodies were used to detect aPDL1, platelets and PMPs. For H&E staining, animals with lung tumours were sacrificed for analysis. The lung tissue sections were stained with H&E following the standard protocol 56 . All H&E staining sections were examined under an optical microscope (Leica DM5500 B).
The local and plasma levels of IL1β, IL6, TNFα and sCD40L were measured using ELISA (eBioscience). For determination of the local (wound) concentration of these cytokines 6 h after platelet injection, the wound tissue was collected and cultured at 37 °C for 24 h. Then 100 μl of medium was removed and frozen at −80 °C for analysis. For the plasma levels, about 6 h after platelet injection, plasma samples were extracted from the mice after treatment and diluted for analysis. All measurements were carried out in triplicate.
The tumours were dissected from the mice and snap frozen in OCT medium. Several 1-μm-thick sections were cut using a cryotome and then mounted on slides. Sections were fixed in ice-cold acetone for 10 min prior to rehydration with PBS. After blocking with BSA (3%), sections were stained with primary antibodies overnight at 4 °C. Following the addition of fluorescently labelled secondary antibodies, the slides were analysed using a confocal microscope (Zeiss LSM 710).
In vivo bioluminescence and imaging
Bioluminescence images were acquired using an IVIS Spectrum imaging system. Living Image software (version 4.4; Perkin Elmer, USA) was used to acquire the data 10 min after intraperitoneal injection of d-luciferin (Thermo Fisher Scientific, Pierce, Cat no. PI88291) in Dulbecco’s PBS (15 mg ml−1) into the animals (10 μl g−1 of body weight). Exposure time for the bioluminescence imaging was 5 min; to optimize reading time, bioluminescence intensity was acquired for 30 min with an exposure time of 1 min using the IVIS system. Regions of interest were quantified as the average radiance (photons s−1 cm−2 sr−1) using the Living Image software.
Tail bleeding assay
Tail bleeding time was determined by removing 3 mm from the tip of the tail and then immediately immersing the tail in PBS at 37 °C. The end point was defined as the time taken for complete cessation of bleeding to occur.
All results are expressed as the mean ± s.d. or the mean ± s.e.m. as indicated. Biological replicates were used in all experiments unless otherwise stated. One-way analysis of variance (ANOVA) and Tukey post-hoc tests were used when more than two groups were compared (multiple comparisons). Survival benefit was determined using a log-rank test. All statistical analyses were performed using the Prism software package (PRISM 5.0; GraphPad Software, USA, 2007). The threshold for statistical significance was P < 0.05. No statistical methods were used to pre-determine the sample size of the experiments.
Source data for the figures in this study are available on figshare at http://dx.doi.org/10.6084/m9.figshare.4231766 (ref. 57 ). The authors declare that all other data supporting the findings of this study are available within the paper and the Supplementary Information.
How to cite this article: Wang, C. et al. In situ activation of platelets with checkpoint inhibitors for post-surgical cancer immunotherapy. Nat. Biomed. Eng. 1, 0011 (2017).
This work was supported by grants from the Alfred P. Sloan Foundation (Sloan Research Fellowship), NC TraCS, the National Institutes of Health (Clinical and Translational Science Award (CTSA, NIH grant 1L1TR001111) to Z.G.) and a pilot grant from the University of North Carolina (UNC) Cancer Center. We acknowledge L. Huang at UNC at Chapel Hill for providing the B16F10-Luc-GFP and 4T1-Luc-GFP cell lines.