Allergen Specific Immunotherapy Partial Mediation Biology

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Background: Allergen specific immunotherapy (SIT), the only disease modifying treatment for allergic diseases, has been used for about a century while its mechanism of action is not completely understood. There is evidence indicating that FOXP3+ regulatory T-cells (Tregs) are induced during SIT in allergic patients. However, a functional role for FOXP3+ Tregs in SIT has not been shown and is difficult to determine in human studies.

Objective: To analyze the kinetics of FOXP3+ Treg induction and to examine whether these cells are required for the beneficial effects of SIT.

Methods: We have previously established a mouse model for SIT in the ovalbumin (OVA)-driven mouse model of allergic asthma, allowing dissection of the relevant mechanisms. In our model, mice were sensitized by OVA/alum, followed by subcutaneous injections of OVA as SIT. We observed a significant induction of FOXP3+ Tregs in spleen and blood? shortly after the SIT injections. Next, we depleted FOXP3+ Tregs after SIT treatment in FOXP3-DTR transgenic mice, followed by inhalation challenges with aerosolized OVA.

Results: Our data show that depleting Tregs after SIT treatment induced a partial but significant (P<0.005) reversal of the suppressive effects of SIT on airway eosinophilia (67%). Moreover, our results reveal that upon depletion of Tregs, SIT fails to significantly suppress allergen induced airway hyperreactivity and serum levels of OVA-specific IgE.

Conclusion: We conclude that FOXP3+ Tregs are partially responsible for the beneficial effects of SIT in our mouse model of asthma.


AHR: airway hyperreactivity

DEREG: depletion regulatory T-cells

DT: diphtheria toxin

i.p.: intraperitoneal

ova: ovalbumin

s.c.: subcutaneous

SIT: specific immunotherapy

Treg: regulatory T-cell


Allergen specific immunotherapy (SIT) is a unique treatment strategy for IgE mediated allergic diseases, that targets the root causes of the disease, leading to long lasting relief of the disease symptoms [1]. The basic principles of SIT were first described by Noon and Freeman at the beginning of 20th century which many of those are still valid today [2,3]. Typically, SIT is performed by repeated monthly exposure to an optimal high dose of the sensitizing allergen for a period of three to five years [4]. Although SIT is mostly effective in allergic rhinitis, allergic conjunctivitis, and venom allergy especially in monosensitized patients [1,5], its efficacy in patients with allergic asthma and multiple sensitizations is controversial and pharmacotherapeutic strategies are often preferred in these patients [6]. Currently, SIT is accompanied by some drawbacks including: need for long term treatment, high risk of anaphylactic reactions and controversy in the treatment outcome. Improving SIT by overcoming the mentioned disadvantages is of particular importance to encourage children to participate in the treatment, since SIT can prevent the progression of respiratory allergies to allergic asthma as well as reducing the chance of developing multiple allergies [7]. SIT improvement is dependent upon an exact and detailed understanding of the mechanisms by which SIT induces tolerance, a knowledge that is lacking up to date.

Although the mechanism of action of SIT is incompletely understood, increased numbers of FOXP3+ cells after SIT has been observed in some clinical studies, while, the functional importance of Tregs in different stages of SIT has not been clearly shown yet. It has been observed that grass pollen SIT increases the frequency of FOXP3+ regulatory T-cells in nasal mucosa [8]. Similarly, expansion of blood FOXP3+CD25hi cells has been observed during venom SIT [9]. In an open-labeled clinical study, it was observed that the levels of induced regulatory T-cells correlate with the outcome of the performed SIT [10]. It is shown that long term depletion of CD4+CD25+T-cells in a mouse model impairs the induction of tolerance [11]. Nonetheless, it is not clear whether regulatory T-cells are critically required at the time of performing SIT. It is also unclear whether maintaining the SIT-induced tolerance is crucially dependent upon regulatory T-cells.

Naturally occurring Tregs (nTregs) constitutively express FOXP3 and high levels of interleukin-2 receptor alpha chain (CD25), folate receptor-4 and Cytotoxic T-Lymphocyte Antigen 4. They are originated in thymus and play indispensable roles in tolerance maintenance against endogenous as well as harmless environmental antigens [12]. Under special circumstances, conventional naïve T-cells in the periphery can also acquire FOXP3 and exhibit regulatory activities, which are equally important for tolerance to self-antigens [12].

Here, we first address one of the most important questions concerning the induction phase of SIT by answering the question, whether nTregs are required for the induction of SIT by depleting these cells just before performing SIT in our previously established mouse model of SIT. Second, we set out to test whether SIT increases and number of FOXP3+T-cells and to find the location where FOXP3+ T-cell are increased upon SIT. Finally we examined whether beneficial effects of SIT are dependent on FOXP3+T-cells using depleting antibodies and FOXP3-DTR transgenic mice. We show that depletion of nTregs during SIT does not abrogate the beneficial effects of SIT. We observed that the number of FOXP3+ CD4+ T-cells is transiently increased in the spleen upon SIT, however, we show that the beneficial effects of SIT are only partially mediated by FOXP3+ T-cells.


Specific pathogen free (according to the Federation of European Laboratory Animal Science Associations (FELASA) guidelines) 6-8-week-old female BALB/c mice were purchased from Charles River laboratories (L'Arbresle, France) and were kept under SPF condition. All animal experiments were performed in accordance with the guidelines of the institutional animal care and use committee of the University of Groningen. Experiment using "depletion of regulatory T-cells" (DEREG) mice was first performed in the animal facility of University of Ghent, Belgium. A follow up experiment using DEREG mice was performed in animal facility of Twincore institute in Hanover, Germany. Non-transgenic littermates of DEREG mice were used as controls.

Experimental allergic asthma, SIT

A shortened protocol was used to induce experimental allergic asthma and to perform SIT as described elsewhere (244). In Brief, mice received 2 intraperitoneal (i.p.) injections of 10 µg ovalbumin (ova; Seikagaku Kogyo, Tokyo, Japan) + 2.25 mg alum (Pierce, Rockford, IL, USA) in 100 µL of pyrogen-free saline on days 0 and 7. Two weeks after the last i.p., they received three subcutaneous (s.c.) injections of 1 mg ova in 200 µl pyrogen free saline on alternate days. Ten days after the 3rd s.c. injections mice were exposed to ova inhalation challenges 3 times every third day. Airway responsiveness to increasing doses of methacholine was measure 24 hours after the last challenge then mice were dissected and samples were taken.

Treg depleting protocol

As shown in figure 1, in experiment B to deplete Tregs before SIT, either anti-CD25 (500 µg/mouse), anti-FR4 (25 µg/mouse) or rat IgG (500 µg/mouse) were administered intraperitoneally one day before every s.c. ova SIT injections. In experiment C, to deplete induced Tregs after SIT, either anti-FR4 (25 µg/mouse) or rat IgG (25 µg/mouse) were injected once intraperitoneally one day prior to the first ova challenge. As illustrated in figure 1, to specifically deplete FOXP3+ regulatory T-cells after SIT, unnicked diphtheria toxin (DT, 100 ug/mouse, Merck, Darmstadt, Germany) to transgenic DEREG mice and their non-transgenic littermates (as control for depletion) one day prior to the first ova challenge.

Evaluation of airway responsiveness

In the last DEREG experiment airway responsiveness to increasing doses of methacholine was evaluated by directly measuring airway resistance as explained elsewhere [13]. Briefly, tracheotomized (20-gauge intravenous: i.v. cannula; Becton Dickinson, Alphen a/d Rijn, The Netherlands), paralyzed (i.v. injection of Pancuronium bromide: Pavulon, 50 µg/Kg Merck Sharp & Dohme, NJ, USA) mice were attached to a computer-controlled small-animal ventilator (Flexivent; SCIREQ, Montreal, Quebec, Canada) under general anesthesia (by i.p. injection of ketamine 100 mg/kg; Pfizer, New York, NY and medetomidine 1 mg/kg; Pfizer). Mice were then ventilated at a breeding frequency of 300 breaths/min and a tidal volume of 10 mL/kg. Tidal volume was pressure limited at 300 mm H2O. An i.v. cannula was placed through the jugular vein for methacholine administration. Thereafter, mice Resistance in response to intravenous administration of increasing doses of methacholine (acetyl-b-methylcholine chloride, Sigma-Aldrich) was calculated from the pressure response to a 2-second pseudorandom pressure wave. In the experiments using antibodies airway responsiveness to inhaled methacholine (Sigma-aldrich) was measured twice (before and after ovalbumin inhalation challenges) in conscious, unrestrained mice using barometric whole-body plethysmography by recording respiratory pressure curves (Buxco; EMKA Technologies) as described in detail previously [14].

Determination of serum levels of ovalbumin-specific IgE

After measuring airway responsiveness, blood was taken, serum samples were prepared and stored at a temperature of -80°C until further analysis. Serum levels of ovalbumin-specific IgE were determined by enzyme-linked immunosorbent assay (ELISA) as described previously and results are expressed as EU/ml.

Analyses of the BAL fluid

Bronchoalveolar lavage (BAL) was performed as explained previously. In brief, animals were lavaged five times through the tracheal cannula with 1-ml aliquots of saline containing a cocktail of protease inhibitors (complete mini tablet (Roche Diagnostics) and 1% bovine serum albumin (BSA: Sigma-aldrich). BAL cells were counted, and cells types were indentified using flow cytometry and according to a published protocol [15]. Flow cytometry data were analyzed using FlowJo (Treestar, RO, USA) and CD3e-,CD19-,CD11c-,CCR3+ cells were considered as Eosinophils.

Preparation of lung tissue for cytokine measurement

Cardiac lobe of lung was taken, homogenized and used to measure cytokine levels as described previously. Concisely, lung tissue was homogenized in 20% (w/v) luminex buffer (50 mM Tris-HCl, 150 mM NaCl, 0.002% Tween 20, and protease inhibitor, pH 7.5) on ice. Subsequently, supernatants were collected for cytokine measurement after spinning the lung tissue homogenates for 10 min at 12,000 x g.

Measurement of cytokines

IL-4, IL-5, and IL-13 in the lung tissue were determined by a commercially available ELISA kit according to the manufacturer's instructions (BD Pharmingen, NJ, USA). The detection limits were 32 pg/ml for IL-5 and 15 pg/ml for IL-4 and IL-13.

Isolation of LNs

To analyze the expression of FOXP3+ regulatory T-cells lymph nodes from different locations were taken after dissecting the mice. Head draining (mandibular, accessory mandibular and superficial parotid) lymph nodes in one pool, forelimb draining (proper axillary and accessory axillary) lymph nodes in a different pool were collected together to obtain the sufficient number for analyzing. A sample size of 6-8 animals was used in each group. Single cells suspensions were made by gently force filtering lymph nodes through 70-µm cell strainers (BD Falcon™, NJ, USA) using 1 ml syringe plunger. Red blood cells were destroyed by keeping the cells in ammonium chloride-containing lysis buffer for 2 minutes at room temperature. Cells were washed 2 times with ice-cold phosphate buffered saline + 1% BSA (Sigma-aldrich) and were analyzed by flow cytometry.

Flow cytometry and antibodies

Hybridoma cells (clone TH6) secreting anti-FR4 antibody were kindly provided by Dr.Shimon Sakaguchi (Department of Experimental Pathology, Institute for Frontier Medical Sciences, Kyoto University). Anti-CD25 (clone PC61) depleting antibody was provided by Mr.L. Boon. PerCp labeled anti-CD4 (clone RM4-5) was purchased from BD Bioscience (NJ, USA). PE-labeled anti-mouse FOXP3 (clone FJK-16s), eFluor450-labeled anti mouse CD25 (clone eBio3C7) were purchased from eBioscience (CA, USA). For flow cytometry, single cells were washed with FACS buffer (PBS + 1%BSA + 0.1% NaN3) then were incubated with antibody mixtures for 30 minutes on ice. Thereafter, cells were washed 3 times with FACS buffer. Flow cytometry was performed using LSR-II (BD bioscience, NJ, USA) and data were analyzed by FlowJo (Treestar, OR,USA).

Statistical analysis

Data are expressed as mean  SEM. The airway resistance curves to methacholine were statistically analyzed using a general linear model of repeated measurements. All the other data were compared using student's t-test. A P-value of less than 0.05 was considered significant.

ResultsDepleting Tregs during induction phase partially reverses AHR suppression

In our model, inhalation challenges with aerosolized ova increases airway reactivity to increasing doses of methacholine (airway hyperreactivity), recruit eosinophils to the lungs and airways, and increase ova-specific IgE levels in previously sensitized mice. Two weeks prior to inhalation challenges, mice receive three subcutaneous injections of either ova as SIT or saline on alternate days. SIT-treated control mice show significantly lower level of airway hyperreactivity, less eosinophil recruitments and lower levels of ova-specific IgE in serum when compared to saline-receiving control mice. To examine whether during induction phase of SIT nTregs are crucially required for the induction of tolerance, SIT-treated and asthmatic mice received either anti-CD25 depleting (PC61), anti-FR4 (TH6) depleting, or rat control immunoglobulin 24 hours before each SIT/placebo injection (Figure 1). Administration of anti-CD25 efficiently depleted more than 90% and anti-FR4 depleted about 75% of FOXP3+CD25+ Tregs in the spleen and blood of mice 24 hours after antibody injection (data not shown). The effects of each injection of antibodies last for at least 48 hours. None of the used antibodies affected the basic manifestation of allergic asthma in placebo-treated control groups in this model (Figures 2A-2E). However, anti-CD25 antibody increased IL-4 levels in lung tissue in anti-CD25-treated placebo-receiving asthmatic mice as compared to rat-IgG-treated placebo-receiving control mice (IL-4: 750 in anti-CD25 treated compared to 447 pg/mg tissue in control immunoglobulin treated group, P<0.05, Figure 2E). Depletion of nTregs using anti-CD25 antibody during SIT partially reversed the suppressive effects of SIT on AHR (Penh at dose 25 mg/ml: 5.37 in SIT+anti-CD25 as compared to 3.36 in control SIT, while 9.07 in placebo control group, Figure 2A). Anti-FR4 antibody treatment did not similarly reverse the effects of SIT on AHR (Figure 2B). SIT in both anti-CD25 and anti-FR4 treated groups suppressed airway eosinophilia and ova-specific IgE levels in serum with the same efficiency as it did in control SIT mice (Figures 2C-2E), so depletion of Tregs failed to reverse the suppressive effects of SIT in our model. The levels of IL-4 in the lung tissue were significantly increased in the anti-CD25 receiving placebo group (P<0.05, IL-4: 868±98 in anti-CD25 compared to 354±100 pg/ml in control placebo group, Figure 2E). This increase is significantly suppressed by SIT (P<0.05, IL-4: 453±60 in anti-CD25 SIT compared to 868±98 pg/ml in anti-CD25 receiving placebo, Figure 2E), while there is no difference between anti-CD25-treated SIT and anti-CD25-treated placebo receiving groups. No statistically significant difference in the levels of IL-4 and IL-5 in the lung tissue has been observed between other groups.

FOXP3+ regulatory T-cells are transiently increased after SIT

Next we addressed the question whether SIT increases the frequency of FOXP3+ CD4+ regulatory T-cells. To this aim, mice were sensitized by two i.p. injections on day 0 and day 7 followed by three s.c. injections of 1mg ova on days 21,23 and 25 (Figure 1). Mice were divided into two groups. A group of mice were dissected on day 26 and the other group was dissected on day 29, spleen, blood, head-draining and forelimb draining lymph nodes, were taken for analyzing the frequency of FOXP3+CD4+ T-cells (24 and 96 hours after the last s.c. injection Figure 1). Interestingly, our data show that SIT significantly increases the number of FOXP3+ cells in spleen at day 1 after the last s.c. injection (FOXP3+ of CD4+ T-cells: 15.5% in SIT treated versus 10.5% in placebo treated mice, Figure 3). The number of FOXP3+ cells in the blood is also increased in SIT treated group as compared to placebo-receiving group (FOXP3+ of CD4+ T-cells: 8% versus 6%). Surprisingly, no difference was observed between SIT-treated and placebo-treated control mice at day 4 after the last s.c. injection. Analyzing the number of FOXP3+ CD4 T-cells reveals no difference between SIT-treated and placebo control in lymph nodes at any time point.

Depletion of regulatory T-cells after SIT completely abrogates the effects of SIT on AHR

To examine whether the beneficial effects of SIT are mediated by Tregs, these cells were depleted using anti-FR4 antibody after SIT and just before inhalation challenges. Mice were sensitized by 2 i.p injections of ova/alum on day 0 and day 7 followed by three s.c. injections of 1mg ova on day 21, 23 and 25. They were three times exposed to ova aerosols every third day starting at day 39 and they were dissected on day 46. One day before the first aerosol challenge on day 38, anti-FR4 antibody (25 µg/mouse) or control rat immunoglobulin was injected. Anti-FR4 administration resulted in more that 75% depletion of CD4+CD25+ cells. Interestingly, Treg depletion using anti-FR4 completely reverses the suppressive effects of SIT on AHR (Penh at dose 25: 9.53 in SIT+anti-FR4 compared to 6.66 in SIT control while placebo + anti-FR4 is 8.97 and placebo control is 9.26 ,P<0.05, Figure 4A). Our data show that Treg depletion by anti-FR4 partially reverses the suppressive effects of SIT on the levels of ova-specific IgE in serum (suppression level: 75.0 % suppression in SIT+anti-FR4 compared to placebo+anti-FR4 versus 87.6% suppression in SIT control compared to placebo control, Figure 4B). Unexpectedly, the level of ova-specific IgE in serum is significantly decreased in anti-FR4-treated placebo-receiving asthmatic mice as compared to rat-IgG-treated placebo-receiving control group (ova-specific IgE: 2010 in anti-FR4 placebo versus 4628 EU/ml in control placebo group P<0.05, Figure 4B). Anti-FR4 administration significantly but partially reverses the effects of SIT on the number of eosinophils in BAL (suppression level: 53.0 % suppression in SIT+anti-FR4 compared to placebo+anti-FR4 versus 86.0% suppression in SIT control compared to placebo control, Figure 4C). The suppressive effects of SIT on IL-4 and IL-5 levels in the lung are completely abrogated in the mice treated with anti-FR4 as compared to those treated with control rat IgG (suppression level for IL-4: 7.5% non-significant suppression in SIT+anti-FR4 compared to placebo+anti-FR4 versus 45.0% significant suppression p<0.01 in SIT control compared to placebo control Figure 4D open bars, suppression level for IL-5: 34.1% non-significant suppression in SIT+anti-FR4 compared to placebo+anti-FR4 versus 54.6% significant suppression p<0.01 in SIT control compared to placebo control, Figure 4D black bars). Anti-FR4 administration results in about 25% decrease in the number of CD4+CD25- cells in the spleen.

Selective depletion of FOXP3+ regulatory T-cells partially reverses the effects of SIT

Since anti-FR4 antibody affected the levels of ova-specific IgE serum as an asthma phenotype in the previous experiment in placebo-treated anti-FR4-receiving group compare to placebo control group and to precisely address the role of FOXP3+ T-cells in mediating the beneficial effects of SIT, transgenic DEREG mice and their non-transgenic littermates were used in our SIT model. SIT was performed according to our optimized protocol and similar to the previous experiments. FOXP3+ Tregs were selectively depleted in transgenic DEREG mice by administration of DT and one day before inhalation challenges (Figure 1). DT administration (1 µg /mouse) in transgenic DEREG mice resulted in more that 90% depletion of CD4+FOXP3+ regulatory T-cells as compared to their non-transgenic littermates (data not shown). Ova challenges significantly increase the airway reactivity to the increasing doses of methacholine both in DEREG transgenic and their non-transgenic littermates as compared to PBS challenges (P<0.05, Figure 5A). As it is shown in figure 5A SIT suppresses AHR in Treg depleted mice with the same efficiency as it does in non-depleted mice. Interestingly, selective depletion of Tregs significantly but partially abrogates the effects of SIT on the levels of ova-specific IgE in serum (suppression level: 84.2 % suppression in SIT compared to placebo in transgenic versus 90.7% suppression in SIT compared to placebo in non-transgenic mice, Figure 5B). Similarly, the effect of SIT on the number of eosinophils in BAL was significantly reversed in Treg depleted mice compared to those without Treg depletion (7.8x106 in Treg depleted compared with 1.9x106 in mice without depletion, P<0.05, Figure 5C). SIT significantly suppresses the levels of IL-4 and IL-5 in the lung tissue in non-transgenic mice. In Treg depleted mice however, this suppressive effect in abolished (P<0.05 in non-transgenic mice, Figure 5D).


In this study we demonstrate that Tregs are dispensable for the induction of SIT in a mouse model of ova-derived allergen specific immunotherapy. We show that FOXP3+ Tregs are transiently induced by SIT, however the beneficial effects of SIT are partially mediated by FOXP3+ Tregs in the used model.

We show that Tregs are not critically required during the induction phase of SIT for the development of an efficient tolerance against ova in a mouse model of SIT. Our results indicate that performing SIT in Treg-depleted and Treg-competent mice induces a similarly efficient tolerance against ova, suggesting that tolerance induction by SIT is independent of Tregs in our mouse model. Our findings partly contradict the study of Boudousquie et al, where they observed tolerance impairment due to a long-term depletion of CD4+CD25+T-cells a mouse model [11]. Although, anti-CD25 was used in that experiment to deplete CD4+CD25+T-cells similar to our approach, it should be noted that they used a different model which differs from ours in the route tolerance induction and its timeframe. Moreover, CD4+CD25+ cells were depleted in that experiment both in the induction and effector phases. Although CD25+FOXP3+ cells were highly depleted in case of usage of either anti-FR4 (about 75%) or anti-CD25 (more than 90%) antibodies. Making a firm conclusion based on the results obtained from antibody depleting experiments is challenging. It could be argued that even low number of Tregs might be sufficient for inducing a highly suppressive SIT.

SIT temporarily increases the frequency of FOXP3+ CD4+ regulatory T-cells in spleen and blood but not in any of the regional draining lymph nodes. We have demonstrated that head and forelimb draining lymph nodes are the major antigen presentation and effector sites. It is only after down-regulation of activation marker, CD69, that proliferating CD4+ T-cells migrate to spleen where the number of FOXP3+ CD4+ T-cells is increased by almost 100%. FOXP3+ CD4+ T-cells in spleen then migrate through the blood which is probably the underlying reason for the increased number of FOXP3+ T-cells in the blood. Inline with our observation, Boudousquie observed a transient induction of FOXP3+ CD4+ T-cells in the lungs after intranasal antigen administration [11]. Given the rate of CD25+CD4+T-cells depletion and high suppressive effects of SIT in experiment A, in the mice receiving either anti-CD25 or anti-FR4, increased number of FOXP3+ cells appears not to be necessary for inducing the suppressive effect of SIT.

We provide evidence indicating that FOXP3+ Tregs are only partially responsible for the suppressive effects of SIT. Selective depletion of FOXP3+T-cells after SIT partially abrogated the suppressive effects of SIT on airway eosinophilia and antigen-specific levels of IgE in serum, suggesting that induction of FOXP3+ cells is not the main mechanism by SIT induces tolerance against the model allergen in our model. It has been previously shown that SIT in our model is completely dependent on IL-10 [16]. Clinical data have also revealed that grass pollen SIT increase the levels of IL-10 in the blood of allergic patients [17] while peptide immunotherapy generates IL-10 dependent tolerance [18]. Bohle and colleagues have observed that sublingual SIT induces antigen-specific IL-10 producing Tregs in the blood of patients [19]. Taken together, partial effects of FOXP3+ T-cells, dependency on IL-10 and long-lasting effects of SIT in our mouse model of SIT and clinical data suggest that a subset of FOXP3 negative Tregs and most likely IL-10 producing Tr-1 cells are responsible for the effects of SIT. Nevertheless, the possibility of clonal deletion of Th2 cells during SIT can not be completely excluded.

In summary we show that Tregs are dispensable during the induction phase of SIT. We show that SIT transiently increases the number of FOXP3+ Tregs in the spleen. We also provided evidence indicating that FOXP3+ cells are partially responsible for the beneficial effects of SIT. It is suggested that either a subtype of FOXP3-Tregs (e.g. Tr-1 cells) or clonal deletion of Th2 cells are the main mechanisms underlying the induction of an efficient SIT.

Article name: Allergen Specific Immunotherapy Partial Mediation Biology essay, research paper, dissertation