Local effector failure in mesothelioma is not mediated by CD4⁺ CD25⁺ T-regulator cells

Mice

Female C57Bl/6J (H-2K^b) mice aged 6–8 weeks were obtained from the Animal Resources Centre (Murdoch, Perth, Australia) and maintained under standard housing conditions. The T-cell receptor (TCR) transgenic mouse line, OT-1, expressing a TCR recognising the H-2K^b restricted dominant OVA_257–264 peptide SIINFEKL (OVAp) 30, was kindly supplied by F. Carbone and W. Heath (University of Melbourne, Melbourne, Australia). GK mice are transgenic for the depleting anti-CD4 antibody (Ab), GK1.5, so have no peripheral CD4 T-cells 31. GK and perforin^−/− mice 32 were bred and housed at the Walter and Eliza Hall Institute (WEHI; Melbourne).

Murine mesothelioma tumour cell line, tumour induction and in vivo growth

AE17 is a MM cell line derived from the peritoneal cavity of C57BL/6J mice injected with asbestos fibres; both AE17 and its OVA transfectant, AE17-sOVA have been previously described 23. On day 0, mice were injected s.c. into the right hind flank with 5×10⁵ tumour cells in 100 μl of PBS, and the rate of tumour growth was measured (mm²) using micro-callipers. All procedures were performed with approval by the University of Western Australia and the Curtin University (both Perth), and the WEHI Animal Experimentation Ethics Committees (AEC). In general, the AEC only permitted tumour growth up to 100 mm²; however, the Curtin University AEC gave conditional short-term approval for an end-point of 150 mm² so that treatment efficacies could be assessed in greater depth.

Harvesting tissues

Tissues were removed and either embedded in Tissue-Tek® O.C.T. compound (Sakura Finetek, Zoeterwoude, the Netherlands) and immediately cryopreserved for immunohistochemistry, or prepared as single cell suspensions by mashing gently between two frosted glass slides in PBS/2% fetal calf serum.

In vivo analysis of antigen presentation (Lyons–Parish assay)

5,6-carboxy-fluorescein-succinimidyl-ester (CFSE; Molecular Probes, Eugene, OR, USA) labelling was performed as previously described 33. Lymph node (LN) cells from TCR transgenic OT-1 mice were incubated with CFSE (5 mM in dimethyl sulfoxide) for 10 min at room temperature. Cells were washed and 10⁷ cells i.v. injected into each recipient mouse. CFSE-labelled cells were analysed by fluorescence-activated cell sorter (FACS) analysis 3 days post-adoptive transfer.

In vivo analysis of CTL function (“in vivo CTL assay”)

Target cells for in vivo evaluation of cytotoxic activity were prepared as described elsewhere 23. Briefly, C57BL/6J LN cell suspensions were red blood cell-lysed, washed and divided into two populations. One population was pulsed with 10⁻⁶ M OVAp for 90 min at 37°C, washed in PBS and labelled with a high concentration (5 μM) of CFSE. Control, uncoated target cells were labelled with a low concentration of CFSE (0.5 μM). Of each population, 10⁷ cells were mixed in 200 μL PBS and i.v. injected into each recipient mouse. Specific in vivo cytotoxicity was determined by collecting the relevant organs from recipient mice 18 h post-injection and the number of cells in each target cell population was determined by flow cytometry. The ratio between the percentages of uncoated versus OVAp-coated (CFSE^low/CFSE^high) was calculated to obtain a numerical value of cytotoxicity. To normalise data allowing inter-experimental comparisons, ratios were calculated between the percentages of peptide coated in control versus tumour-bearing mice.

FACS analysis

Single cell suspensions were stained with a combination of CD4 (RM 4–5), CD8α (53–6.7), CD11c (HC3), CD45R/B220 (RA3-6B2), anti-murine interferon (IFN)-γ (XMG12) CD62L (MEL-14), CD69 (H1.2F3), CD44 (IM7), CD25 (3C7; all from Pharmingen, San Diego, CA, USA), F4/80 (clone CI:A3-1; Caltag), CD4 (RM 4-5; eBioscience, San Diego, CA, USA), CD25 (PC61), FoxP3 (150D; Biolegend, San Diego, CA, USA) and/or phycoerythrin-labelled OVAp-H-2K^b tetramer (kindly provided by A. Brooks, University of Melbourne). Analysis was performed on a FACScan (Becton Dickinson, Mountain View, CA, USA) using CellQuest and FlowJo software or on a FACSCanto II using FACSDiva software (all BD Biosciences, San Jose, CA, USA).

Immunohistochemistry

Frozen sections were fixed in cold ethanol. Endogenous peroxidases, avidin and biotin were blocked using 1% hydrogen peroxide and the Avidin/Biotin blocking kit (Dako, Glostrup, Denmark). For single staining, primary Abs directed against murine CD4 and CD8, CD11c, B220 (clone RA3-6B2; Pharmingen), F4/80, CD31 (clone MEC 13.3; Pharmingen) and isotype controls (rat immunoglobulin (Ig) G2_a, rat IgG2_b and hamster IgG) were consecutively linked to a secondary biotinylated Ab (anti-rat monoclonal Ab; Jackson ImmunoResearch, West Grove, PA, USA), or anti-hamster Ab (Pharmingen), followed by streptavidin-horseradish peroxide (Dako). After washing, one Sigma FAST^TM (D-4168) 3,3-diaminobenzidine tablet and one urea hydrogen tablet (Sigma-Aldrich, St Louis, MO, USA) were added to ddH₂O to serve as a peroxidase substrate (125 μL·section⁻¹) and haematoxylin counterstained to visualise staining. Sections were then dehydrated and mounted with DPX (Ultramount; Scot Scientific, Perth).

For double staining, primary Abs were linked to a peroxidase-conjugated Ab (rabbit anti-rat or anti-hamster; Dako) and detected using 3.3’-diaminobenzidine substrate as a peroxidase substrate. A biotinylated primary Ab linked to streptavidin-alkaline phosphate (Dako) was visualised using an alkaline phosphatase substrate (BCIP/NBT; Vector Laboratories, Burlingame, CA, USA). Sections were mounted in Immunomount (Shandon, Pittsburgh, PA, USA).

In vivo depletion using monoclonal antibodies

For depletion of CD4⁺ or CD8⁺ cells, two doses (150 μg·dose⁻¹) of either YTS-191 or YTS-169 (European Collection of Animal Cell Cultures, Salisbury, UK), respectively, were injected i.p. with 3 doses·week⁻¹ (100–150 μg·dose⁻¹) for 2 weeks and spleens tested (by FACS analysis); CD8⁺ depletions were 95–99% effective, whilst CD4⁺ depletions were 90–95% effective (data not shown). For depletion of CD25⁺ cells, one or two doses (150 μg·dose⁻¹) of PC61 (anti-mouse CD25 monoclonal Ab obtained from the Monoclonal Antibody Facility, Perth) in 100 μL PBS were injected intra-tumourally or peri-tumourally 34. The endotoxin levels in PC61 were <0.1 EU·mL⁻¹ (measured by supplier using an endotoxin detection kit, manufacturer's documentation supplied).

IL-2 and IL-21 immunotherapy

Lyophilised proleukin (recombinant IL-2; Cetus Corporation, Emeryville, CA, USA) was reconstituted in sterile PBS (Sigma-Aldrich) and given intratumourally as previously described 23. The vector pORF/mIL-21 (Invivogen, San Diego, CA, USA) was used for the production of IL-21 in vivo; 20 μg of plasmid in 2 mL of saline was injected hydrodynamically 35.

Statistical analysis

Statistical significance was calculated using GraphPad PRISM (San Diego, CA, USA). An unpaired t-test was used to determine differences between two populations. One-way ANOVA was used to determine differences between more than two populations.

DCs are located in the mesothelioma tumour microenvironment

DCs are highly efficient at capturing and processing in situ antigen for presentation to T-cell after trafficking to dLN. Thus, tumour-infiltrating DCs represent a likely candidate for priming the anti-tumour response. Therefore, the first series of studies examined tumour-infiltrating DCs in progressing tumours that were divided into previously defined MM tumour sizes based upon their responsiveness to IL-2 immunotherapy; i.e. tumours <25 mm² are readily cured after intratumoural IL-2 treatment, whilst 100% of those >25 mm² completely fail to respond 23. Therefore, small tumours are defined as those <25 mm², medium sized tumours are 25–50 mm² and large tumours are 50–100 mm². Note that in accordance with AEC conditions we are unable to grow tumours beyond 100 mm². Tumours were sampled when they were small versus medium sized.

CD11c is a pan DC marker and CD11c⁺ cells were found infiltrating tumours (fig. 1a⇓) and their proportional abundance did not significantly alter during tumour progression. Interestingly, the percentage of CD11c⁺ DCs in dLN in animals bearing medium tumours was significantly higher than DCs levels seen in normal LN controls (fig 1a⇓).

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Fig. 1—

Dendritic cell (DC) presentation of malignant mesothelioma (MM)-antigens to naïve T-cells generates effector cytotoxic lymphocyte (CTLs); tumour and draining lymph nodes (dLN) single cell suspensions prepared from mice bearing small and medium tumours were stained with anti-CD11c for DCs. a) Pooled data are from two experiments (6 mice·group⁻¹) and represented as mean±sem. Tumour: ▒; dLN: ▪. *: p<0.05 comparing tumour dLN with normal lymph node (LN). To analyse MM-antigen presentation to naïve T-cells 5,6-carboxy-fluorescein-succinimidyl-ester (CFSE)-labelled, tumour antigen-specific, OT-I T-cells were adoptively transferred into (b, c) recipient AE17-secreted ovalbumin (sOVA)-bearing mice 3 days prior to analysis. dLN (b) and non-dLNs (c) were harvested from recipient mice, prepared as a single cell suspension and stained for CD8. Fluorescence-activated cell sorter (FACS) analysis was performed by gating on CD8⁺/CFSE⁺ cells. Mice inoculated with AE17 tumour cells were negative controls (d and e). Representative histograms are shown (b–e). In vivo CTL activity was assessed by adoptively transferring differentially-labelled target cells prepared from normal mice representing CFSE^high or OVA_257–264 peptide SINFEKL (OVAp), and CFSE^low control cells into mice bearing AE17-sOVA (f and g) or AE17 (h and i) tumours. dLN and non-dLN were prepared as a single suspension 18 h later and FACS analysed. A reduction in the OVAp peak compared with the control peak represents lysis of the targets. Representative histograms are shown (f–i). CTL activity is shown as the number of cells in the OVAp peak divided by the number of cells in the control peak multiplied by 100. All data within each experiment were normalised compared with nontumour bearing C57BL/6J LN controls. j) Pooled in vivo CTL activity data (9 mice·group⁻¹) from dLN and non-dLN are shown as mean±sem. One way ANOVA was performed comparing small versus medium sized tumours. ▓: non-dLN; ▪: dLN.**: p<0.01.

MM tumour antigens are rapidly transported to dLN for presentation to T-cells

One possible role of tumour-residing DCs is the presentation of tumour antigens to naïve T-cells in dLN. Therefore, we assessed whether tumour antigens are presented in vivo in LN during disease progression using the AE17-sOVA cell line. AE17-sOVA is readily recognised by OVA-specific CTL taken from OT-I mice 23, therefore, antigen presentation was identified by OT-I CD8⁺ T-cell proliferation in vivo. To do this CFSE-labelled OT-I cells were i.v. injected into tumour-bearing mice previously given tumour cells. After 3 days, lymphoid organs were collected and OT-I proliferation was assessed by CFSE dilution.

As expected, antigen presentation was primarily seen in the dLN (fig 1b⇑). Unlike other MM models wherein tumour antigen presentation was restricted to the dLN 13, some limited proliferation was seen in non-dLN (fig 1c⇑), presumably because OVA was secreted. Antigen presentation could be detected even when tumours were small. These results show that tumour antigen presentation occurs early in MM tumour growth.

Tumour-specific CTL are initially generated in dLN and spread systemically

Following this, we assessed whether antigen presentation led to the generation of tumour-specific CTL in LN. OVAp-pulsed CFSE-labelled target cells were adoptively transferred into AE17-sOVA tumour-bearing mice, and lymphoid organs were analysed by FACS 18 h later. OVAp-specific CTL in dLN could be detected in mice with small tumours and their killing activity significantly increased with increased tumour burden (fig 1j⇑). Note that as tumour burden increased OVAp-specific CTL were also found at equivalent levels in non-dLN implying systemic spread of these CTL (fig 1f⇑, g and j). These results show that tumour antigen presentation and the generation of tumour-specific CTL occurs early in MM tumour growth.

Small numbers of CD8⁺ T-cells infiltrate MM tumours

Next we wanted to address whether T-cells generated in the dLN migrated into the MM tumour microenvironment. Initially, this was assessed using FACS analysis (fig. 2a⇓) of tumours disaggregated to single cells and immunohistochemistry on frozen tumour sections (data not shown); both were stained with anti-CD8 antibodies. Small percentages (3.4–5.2%) of CD8⁺ cells were present within MM tumours (fig. 2a⇓). Interestingly, the percentage of CD8⁺ cells within LN increased in small tumour-bearing mice but fell to normal levels as MM tumours progressed.

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Fig. 2—

CD8⁺ cells penetrate the malignant mesothelioma tumour microenvironment. Single cell suspension of small or medium tumours and their lymph nodes (LNs) were stained for CD8. a) Pooled data from three experiments (9 mice·group⁻¹) are shown as mean±sem. ▪: tumour; ░: draining LN (dLN); ▒: non-dLN. ***: p<0.001 comparing dLN and non-dLN with normal LN. Frozen tumour sections were double stained for b, d and e) CD8⁺ cells (blue) and c, d and e) CD31 to detect blood vessels (brown). b) Rat immunoglobulin (Ig) G2a and c) rat IgG2b are isotype controls. These experiments were performed twice (6 mice·group⁻¹) and representative photos at 200 × magnification are shown. e) The inset shows a CD8⁺ cell associated with a tumour blood vessel and a CD8⁺ cell within the tumour matrix, indicated by arrows.

Double staining of tumour blood vessels (using anti-CD31 or platelet/endothelial cell adhesion molecule-1) with CD8 showed that CD8⁺ cells could be found outside blood vessels and within the tumour itself (figs. 2b–e⇑). Thus, some CD8⁺ T-cells appear to be able to undergo diapedesis through tumour-associated blood vessels to penetrate the tumour matrix.

CD8⁺ T-cells in mesothelioma tumours are activated

The tumour microenvironment was also assessed for signs of local antigen presentation (from the same animal shown in fig. 1b–e⇑) after adoptive transfer of CFSE-labelled OT-1 cells. Very few OT-1 cells had penetrated the tumours 3 days after transfer, however, those that had appeared to have proliferated (fig. 3b⇓), unlike the same OT-I cells transferred into mice given OVA in incomplete Freunds adjuvant (fig. 3a⇓), there were no remaining parental cells. However, we could not exclude the possibility that they represent cells that proliferated elsewhere and then migrated into the tumour. Similarly, the tumour microenvironment was assessed for signs of local CTL activity from the same mice used for fig. 1e–j⇑. While there was clear CTL activity in the dLN (fig. 3c⇓), the few target cells that could be detected in the tumour microenvironment showed no sign of loss of OVAp-pulsed target cells (fig. 3d⇓) suggesting that intratumoural CTL had lost the killing ability they acquired in the dLN.

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Fig. 3—

Malignant mesothelioma-infiltrating CD8⁺ T-cells are activated but lose cytotoxic lymphocyte (CTL) activity; tumours were also harvested from AE17-secreted ovalbumin (sOVA)-bearing mice given 5,6-carboxy-fluorescein-succinimidyl-ester (CFSE)-labelled, tumour antigen specific, OT-I T-cells (as per fig. 1b⇑ and c). a) The few OT-I cells that penetrated the tumour expressed low levels of CFSE similar to proliferating cells seen in mice given OVA in incomplete Freunds adjuvant draining lymph node (dLN), indicative of proliferation. However, the few CFSE^high (OVA_257–264 peptide SINFEKL) and CFSE^low target cells harvested from the tumours of d) AE17-sOVA-bearing mice (as per fig. 1f⇑ and g) showed no in vivo CTL activity relative to c) the CTL activity in dLN of the same mouse. In separate experiments, tumours and dLN harvested from AE17-bearing mice were double stained for e) CD8 and CD69, f and g) interferon (IFN)-γ, h) CD25 and i) CD44. Fluorescence-activated cell sorter analysis was performed by gating on CD8⁺ cells. Representative histograms are shown with dLN and tumour overlaid for e) CD69, f) IFN-γ analysis in the dLN and g) tumour. e) ––––: tumour; ······: dLN. f and g) –––––: IFN-γ; ········: rat immunoglobulin G1 isotype control. These experiments were repeated twice (6 mice·group⁻¹). Pooled data from two experiments (6 mice·group⁻¹) is shown for h) CD25 and i) CD44 as mean±sem percentage of CD8 cells. ▪: normal LN; ▓: AE17; □: AE17-sOVA. **: p<0.01, comparing dLN with normal LN.

The activation status of MM-infiltrating CD8⁺ T-cells was also assessed using a range of markers including the early activation marker CD69 (fig. 3e⇑), IFN-γ (fig. 3f⇑ and g) and the peripheral LN homing marker, CD62L (data not shown), CD25 (fig. 3h⇑) and the late activation marker, CD44 (fig. 3i⇑). As expected, CD62L was downregulated on tumour-infiltrating CD8⁺ T-cells relative to those from LN. All other markers were upregulated on CD8⁺ T-cells in small MM tumours relative to their expression on CD8⁺ T-cells in normal LN or in tumour-dLN. In particular, CD25 and CD44 were highly co-expressed by tumour-infiltrating CD8⁺ T-cells. CD8⁺ T-cells in tumour-dLN expressed higher levels of CD25, but not CD44, than their healthy counterparts (fig. 3h⇑ and i).

Tumour-specific CD8⁺ T-cells accumulate in MM tumours

The major histocompatibility complex class I SIINFEKL tetramer was used to identify endogenous tumour-specific CD8⁺ T-cells. Tetramer⁺CD8⁺ T-cells were detected in low proportions (<1.6% of total CD8 population) in dLN throughout tumour growth (fig. 4b⇓ and e). In contrast, much higher levels (up to 15.7% of CD8⁺ T-cells were tetramer⁺) were seen in MM tumours (fig. 4d⇓ and f). These tetramer⁺ cells were activated, as shown by CD44 and CD69 expression (fig. 4g–i⇓). Taken together, these data clearly show that, after tumour antigen is presented to naïve T-cells in dLN, fully functional tumour-specific CTL are generated which home to the tumour. However, once in the tumour site their effector function appears compromised.

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Fig. 4—

Tumour-specific CD8⁺ cells in malignant mesothelioma tumours are activated. a and b) Draining lymph node (dLN) and c and d) tumours were harvested from a and c) AE17 and b and d) AE17-secreted ovalbumin (sOVA) tumour-bearing mice and fluorescence-activated cell sorter analysed for co-expression of the OVA_257–264 peptide SIINFEKL (OVAp) tetramer and CD8 (a–f), as well as g) an isotype control, h) CD44 or i) CD69. Flow cytometric analysis was performed by gating on CD8⁺tetramer⁺ cells. a–d) Representative dot plots and g–i) histograms are shown. Pooled data from two experiments (6 mice·group⁻¹) are shown as mean±sem percent of CD8 cells that are tetramer⁺ in the e) dLN and f) tumour. ░ AE17; □: AE17-sOVA. ***: p<0.001 comparing small and medium tumours.

Activated CD4⁺ T-cells are also located in the MM tumour microenvironment

The presence of activated CD8⁺ T-cells within MM tumours does not provide much benefit to the host as the disease continues to progress implying local immunosuppression. One suppressive cell type that has been recently identified is the CD4⁺ regulatory T-cell. Thus, we examined CD4⁺ T-cells in MM tumours versus those in tumour-dLN and non-dLN, and healthy LN. LN taken from MM-bearing hosts, regardless of tumour burden, contained significantly more CD4⁺ T-cells (27.9±1.0% to 30.4±1.3%) than normal LN (20.6±0.7%; fig. 5a⇓). However, the ratio of CD8:CD4 in tumour-dLN was 1.23:1; thus CD8⁺ T-cells outnumbered CD4⁺ T-cells. CD4⁺ cells also infiltrated MM tumours in small numbers. The percentage of CD4⁺ cells (ranging from 4.7% to 6.2%) within tumours did not change with tumour burden and a ratio of 1:1 of CD8:CD4 was seen in the tumour bed. Similar to CD8⁺ T-cells, MM-infiltrating CD4⁺ T-cells were also activated as they expressed CD69, CD44 and secreted IFN-γ (fig. 5c–j⇓).

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Fig. 5—

Malignant mesothelioma-infiltrating CD4⁺ T-cells are activated. Tumours and draining lymph nodes (dLN) were harvested from AE17-bearing mice and stained for CD4 expression and fluorescence-activated cell sorter (FACS) analysed. These experiments were repeated twice (6 mice·group⁻¹). a) pooled data are shown as mean±sem percent of CD4⁺ cells. *: p<0.05 and **: p<0.01 comparing dLN (░) and non-dLN (▒) with normal LN (□). ▪: tumour. Cells were also double stained for CD4 and c and g) isotype controls, or d and h) CD69, e and i) CD44 as well as f and j ) interferon (IFN)-γ. FACS analysis was performed by gating on CD4⁺ cells and representative FACS histograms are shown (c–j). f and j) ––––: IFN-γ; ······: isotype control. CD4⁺CD25⁺ co-expression was also examined in normal LN, dLN and tumours and b) pooled data from two experiments with (6 mice·group⁻¹) are shown as the percent of CD4⁺ that are CD25⁺ ±sem.

Expression of the α-chain of IL-2R (CD25) was also examined on CD4⁺ T-cells in MM-bearing hosts. CD25 expression can be indicative of activation; however, CD25 co-expression with the nuclear transcription factor molecule, FoxP3, is recognised to be a marker of T-regulatory (Treg) cells. CD25 expression levels were not increased on CD4⁺ T-cells in MM-draining LN relative to healthy controls (fig. 5b⇑). In contrast, increased expression levels of CD25 was seen on MM-infiltrating CD4⁺ T-cells (>20% of CD4⁺ cells).

CD4⁺ cells are more likely to function as effector and not suppressor cells

Depletion studies were conducted to identify the function that CD4⁺ T-cells exert on progressing tumours. As the depleting Abs are rat Abs they could only be used for 2 weeks before destruction by an anti-rat immune response. Transient removal of CD4⁺ T-cells did not have any impact upon tumour growth rate (fig. 6a⇓). Use of mice that never have a mature CD4⁺ T-cell compartment (GK mice) resulted in a faster tumour growth rate that was significantly different to that seen in immunologically intact wild-type mice (fig. 6b⇓), suggestive of a CD4 effector, and not regulator, phenotype. Thus, global removal of CD4⁺ cells appears to remove an effector cell.

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Fig. 6—

The majority of CD4⁺ cells are effector cells. The role of CD4⁺ cells was determined by a) antibody depletion (○: CD4 depletion; ▪: no depletion), and b) by the use of GK mice (mice that never have a mature CD4⁺ T-cell compartment). Pooled data are shown as mean±sem tumour size from 5 mice·group⁻¹. b) Only GK mice were significantly different to C57BL/6 mice. •: C57BL/6 mice; □: GK CD4-mice. c) To address a possible role for splenic cells, naïve C57BL/6 mice were the recipients of adoptively transferred, unfractionated spleen cells prepared from unmanipulated, healthy mice and from mice with progressing AE17 tumours. All mice, plus tumour growth controls (no transfer), were challenged with AE17 tumour cells 3 weeks later. c) Pooled data from one of two experiments (8 mice·group⁻¹) are shown as mean±sem. ▵: no transfer; ♦: healthy splenocytes; ▴: tumour splenocytes. d) Tumour and draining lymph node (dLN) from AE17 tumour-bearing mice were fluorescence-activated cell sorter analysed for CD4⁺FoxP3⁺ co-expression in normal lymph node (▪), tumour dLN and tumours. Pooled data are shown after gating on CD4⁺ cells from 12 mice·group⁻¹. ░: AE17 mice.

Splenic Treg cells do not ablate protective responses

The spleens of tumour-bearing mice may contain Treg cells. Thus, to gain insights into their role, splenocytes prepared from healthy mice or from tumour-bearing mice were transferred into recipient mice which were then challenged with AE17 tumour cells 3 weeks later. Recipients of splenocytes from tumour-bearing mice were completely protected (fig. 6c⇑), whilst those from unburdened healthy mice were unable to prevent tumour growth. These data show that endogenously-generated, tumour antigen-specific splenic T-cells provide significant levels of protection when transferred into a naïve host, and that co-located suppressor cells do not interfere with this process. Yet the same cells cannot protect the host within which they were induced, as their tumours continued to progress. These data suggest that there is site-specific regulatory activity within the tumour bed.

MM-infiltrating CD4⁺ Treg cells do not exert potent suppressor activity

Analysis of FoxP3 expression on CD4⁺ T-cells (independent of CD25), revealed putative Treg cells within the MM microenvironment (>18%; fig. 6d⇑). There was no proportional increase of FoxP3⁺CD4⁺ T-cells in tumour-dLN relative to normal LN.

Treg cell depletion does not abrogate suppression of anti-MM immune responses

The above data suggested that the site of suppressive Treg cells function may be the tumour microenvironment. Therefore, we wanted to deplete Treg cells in situ using an anti-CD25 monoclonal Ab (PC61), as depletion of CD4⁺ Treg cells by this Ab had been reported previously in several murine cancer models 36, 37 including MM 18, 34. FACS analysis showed that a single intra-tumoural injection of PC61 not only removed CD4⁺CD25⁺FoxP3⁺ cells from the tumour microenvironment (fig. 7a⇓, d, g and j) but also from all organs examined including LN, spleen (fig. 7b⇓, c, e, f, h, i, k and l) and bone marrow for >12 days (fig. 7m⇓). Also note that <30% of tumour infiltrating CD4⁺CD25⁺ cells were FoxP3⁺ (Fig. 6d) suggesting that the remaining CD4⁺CD25⁺ cells may be activated effector cells; these cells were also depleted. Despite the global removal of CD4⁺CD25⁺ cells, MM tumours continued to progress at exactly the same rate as undepleted controls (fig. 7n⇓).

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Fig. 7—

CD25⁺CD4⁺ T-regulatory (Treg) cells do not interfere with endogenous anti-malignant mesothelioma immunity. The role of CD4⁺CD25⁺ suppressor cells was determined by depletion using the anti-CD25 antibody (PC61). A single intra-tumoural injection of PC61 depleted CD4⁺CD25⁺FoxP3⁺ from the tumour (a, d, g and j), draining lymph node (b, e, h and k), spleen (c, f, i and l) and bone marrow for 10 days (m). Representative dot plots 4 or 6 days after PC61 injection show CD4⁺CD25⁺ (a–f) gated on lymphocytes, and FoxP3⁺CD25⁺ gated on CD4⁺CD25⁺ (g–l). a–c) and g–i) not depleted; d–f) and j–l) PC61 depleted. m) ♦: tumour; ▾: bone marrow; ▵ lymph node; □: spleen. Data are from one of three experiments in which the organs from 5 mice·group⁻¹ were pooled for analysis. n) Mice were given one (▴) or two (▿) PC61 injections (arrows) and tumour growth was monitored (three experiments with 5 mice·group⁻¹); data shown as mean±sem. □: no depletion.

Treg cell depletion does not improve immunotherapy

We have previously shown that intratumoural IL-2 immunotherapy is effective at curing small MM tumours 23 and hypothesised that depletion of Tregs may remove a population of cells that function as an IL-2 sink on account of their CD25 expression levels. Thus, removal of Tregs should improve effector cell responses. Instead, in vivo depletion of CD25⁺ using PC61 completely disarmed IL-2 mediated effector function (fig. 8a⇓). As it has been previously shown that IL-2 driven anti-tumour immunity is T-cell mediated 23 we conclude that we removed effector and not regulator cells.

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Fig. 8—

CD25-targeted T-regulator (Treg) cells depletion may confound cytokine-based immunotherapy. a) A single PC61 injection was given 1 day prior to intratumoural interleukin (IL)-2 injections given every 2nd day (two experiments with 5 mice·group⁻¹). b) PC61 injections were given 1 day before each of two hydrodynamic injections of pORF/mIL-21 that were 6 days apart (one experiment with 5 mice·group⁻¹), and tumour growth monitored. ▾: PBS no depletion; ⋄: Treg depletion; •: Il-2 no depletion; □: IL-2 Treg depletion; ▴: saline no depletion; ♦: saline Treg depletion; ○: IL-21 no depletion; ▿: IL-21 Treg depletion; ▓: treatment duration. Arrows: Treg depletion. Data shown as mean±sem.

We also tested the role of Tregs when faced with IL-21 driven anti-tumour immunity. In this case, the removal of CD25⁺ cells was a null event with no evidence of interference with effector or regulator cells (fig. 8b⇑). Taken together, these data suggest either a weak role for Tregs as anti-tumoural weapons, and/or that attempting their removal via CD25 targeting removes effectors as well as regulators.