Induction of respiratory immune responses in the chicken; implications for development of mucosal avian influenza virus vaccines

Abstract

The risk and the size of an outbreak of avian influenza virus (AIV) could be restricted by vaccination of poultry. A vaccine used for rapid intervention during an AIV outbreak should be safe, highly effective after a single administration and suitable for mass application. In the case of AIV, aerosol vaccination using live virus is not desirable because of its zoonotic potential and because of the risk for virus reassortment. The rational design of novel mucosal-inactivated vaccines against AIV requires a comprehensive knowledge of the structure and function of the lung-associated immune system in birds in order to target vaccines appropriately and to design efficient mucosal adjuvants. This review addresses our current understanding of the induction of respiratory immune responses in the chicken. Furthermore, possible mucosal vaccination strategies for AIV are highlighted.

Keywords:

• avian influenza virus
• chicken
• poultry
• mucosal vaccination
• adjuvant
• respiratory tract
• antigen uptake
• humoral immunity
• haemagglutinin
• neuraminidase
• BALT

1. Introduction

Several important poultry pathogens, including avian influenza virus (AIV), infectious bronchitis virus (IBV) and Newcastle disease virus (NDV), enter the host through the mucosal tissues of the respiratory tract (RT) (Villegas 1998) and subsequently disseminate towards other organs in the body. Therefore, animal health significantly depends on the control of infection already in the lung tissue by the RT immune system. The rational design of novel mucosal vaccines against AIV requires a comprehensive knowledge of the structure and function of the lung-associated immune system in birds in order to target vaccines appropriately and to design efficient mucosal adjuvants. At the moment, there is very limited knowledge of the lung-associated immune system in the chicken and in poultry, in general, which might be a consequence of the unique and complex anatomy and function of the avian lung.

2. Avian influenza virus

Influenza virus is a member of the orthomyxoviruses, a family of single-stranded (ss)-RNA viruses. Three types of influenza virus exist: A, B and C. Aquatic birds are the natural reservoir for influenza A (Baigent and McCauley 2003; Muramoto et al. 2006). However, influenza A viruses can infect a wide range of mammalian and avian species. Influenza A viruses are characterized by their two surface antigens (Ag), haemagglutinin (HA; H1–16) and neuraminidase (NA; N1–9). All HA and NA subtypes have been isolated from avian species, however H3, H5, H7 and H9 infections are most common in domestic poultry (Swayne 2008).

Influenza A enters the host cell by binding of HA to sialic acids (SA) attached to galactose (Gal; SAα-2,3Gal or SAα-2,6Gal-linkages) (Gambaryan et al. 1995). Human influenza viruses prefer SAα-2,6Gal-linkages, whereas avian viruses prefer SAα-2,3Gal (Baigent and McCauley 2003). Both SAα-2,3Gal- and SAα-2,6Gal-linked receptors are widely expressed within different tissues, including the RT of poultry (Kuchipudi et al. 2009; Pillai and Lee 2010; Yu et al. 2011). This suggests that poultry can be infected with a wide range of influenza viruses and also probably act as a ‘mixing vessel’.

Following fusion, influenza virus mainly enters the host cell via not only clathrin-dependent mechanisms, but also clathrin-independent and caveolin-independent mechanisms (Lakadamyali et al. 2004), with the characteristics of macropinocytosis (de Vries et al. 2011). Once internalized, these pathways are all equally efficient for virus fusion (Lakadamyali et al. 2004). For productive infections, influenza viruses need to have their HA0 cleaved into HA1 and HA2 segments. Low-pathogenic (LP) AIV HA proteins are cleaved by trypsin-like enzymes located primarily in the RT or produced by some types of bacteria. Highly-pathogenic (HP) AIV possess multibasic cleavage sites and can therefore be cleaved by multiple enzymes (Swayne 2008), thereby facilitating systemic replication (Webster and Rott 1987; Munster et al. 2010).

In chicken, infection with HPAIV causes severe depletion, haemorrhages, oedema, cutaneous ischaemia and mortality rates that can reach up to 100% (Muramoto et al. 2006). The H5N1 HPAIV has had a great impact on the poultry industry in recent years. Moreover, it is a zoonotic infection, with high mortality rates after transmission to humans. In the Netherlands, the last HPAIV outbreak was an H7N7 outbreak in 2003. Although control measures were implemented, 255 poultry flocks became infected at that time and 1381 commercial flocks were depopulated (Stegeman et al. 2004; Elbers et al. 2005). In infected poultry flocks, clinical signs included increased mortality, swollen head and wattles, respiratory problems and decreased egg production (Elbers et al. 2005). During the epidemic, 89 people involved in handling of infected poultry became infected, resulting in one death (Fouchier et al. 2004).

Not only HPAIV infections pose a threat to the poultry industry and to man, but LPAIV strains as well. LPAIV strains are most prevalent and can cause significant economic losses. Furthermore, they can contribute to, or mutate into more pathogenic strains (Marcus et al. 2007). LPAIV infections can cause subclinical to mild symptoms, primarily in the RT. Mortality rates are low, but can significantly increase in the case of secondary infections (de Wit et al. 2004; Swayne 2008).

As AIV enters the host via the RT and from there spreads towards other organs, a vaccine should already halt infection at the mucosal tissues of the RT, the portal of entry. For the design of new vaccines, it is important to gain knowledge on how the RT innate immune system handles viral (vaccine) antigens. Knowledge on early immune responses after infection or vaccination is crucial for the rational design of novel mucosal vaccines. Furthermore, it is important to know if uptake of a vaccine antigen locally in the lung results in induction of both local and systemic humoral responses.

3. Immune system of the avian RT

3.1. Morphology of the avian lung

The chicken RT differs significantly from mammalian RT in both morphology and airflow as well as immunologically. The mammalian lung has bidirectional airflow, whereas the avian lung has a unidirectional airflow (Figure 1A) (Reese et al. 2006). Chicken lungs are located dorsally that the ribs incise the lungs (Duncker 1972). The lungs are small when compared to mammals, and do not significantly change in volume during respiration. Nine air sacs are present, which function in ventilation, but do not contribute to the gas exchange (Figure 1B). These adaptations of the avian lung facilitate rapid gas exchange needed for sustained flight (Farmer 2006; Maina 2006). Furthermore, air sacs could provide birds with the stability and agility needed in flight (Farmer 2006).

Figure 1. Anatomy of the avian lung. A: Schematic drawing of air flow in the avian lung during inspiration (left picture) and expiration (right picture) (adapted from Kothlow and Kaspers 2008, with permission from Elsevier). B: Respiratory system of chickens (Clav. AS = clavicular air sac; Cran. Th. AS = cranial thoracic air sac; Caud. Th. AS = caudal thoracic air sac; Abd. AS = abdominal air sac). During inhalation, the air sacs expand and air is drawn from the outside through the trachea and primary bronchi, partly towards the caudal air sacs and partly towards the paleopulmonic parabronchi (Fedde 1998).

Birds can breathe through their nares or mouth and air first arrives in the oronasal cavity. The oronasal cavity is separated from the trachea by the larynx, which opens into the trachea via the glottis. At the syrinx, the trachea bifurcates into two primary bronchi (Powell 2000). The primary bronchus enters the lung ventrally and passes to the dorsolateral surface, ending up in the abdominal air sacs (Duncker 1972). The secondary bronchi originate from the primary bronchus. Gas exchange occurs at the tertiary bronchi. The blood–gas barrier of the avian lung is much thinner than that of mammals with the same body mass, while the respiratory surface is greater. These features are thought to predispose birds to pulmonary injury (Reese et al. 2006).

The lymphoid tissue of the RT can roughly be divided into an upper and a lower part. The lymphoid tissue of the upper part of the RT includes paraocular (including the Harderian gland (HG)) and nasal lymphoid structures and lymphoid accumulations in the pharynx and larynx. The avian lung exhibits both highly organized lymphoid structures and diffusely-distributed lymphoid and myeloid cells. The organized bronchus-associated lymphoid tissue (BALT) structures develop at the junctions of the primary bronchus and the caudal secondary bronchi (Figure 2) (Bienenstock et al. 1973; Van Alstine and Arp 1988). In contrast to the mammalian lung, where BALT is inducible, it is constitutively present in the chicken lung (Bienenstock et al. 1973). It is believed that BALT is a normal lymphoid structure in poultry, compensating for the lack of a lymph node system (Fagerland and Arp 1993a).

Figure 2. Schematic drawing of a chicken lung indicating locations of BALT. PB = primary bronchus; 1 = medioventral secondary bronchi; 2 = mediodorsal secondary bronchi; 3 = lateroventral secondary bronchi; A–D = openings into the air sacs. BALT nodules are indicated as orange dots (Reese et al. 2006).

BALT development follows a similar time course in specified pathogen-free (SPF) and conventional chickens, though slight differences are observed with regards to germinal centre (GC) development (Fagerland and Arp 1993a). The development is also similar in broiler and layer chickens and in turkeys. In one-day-old chickens, no immunoglobulin (Ig)-producing cells of any isotype are present (Fagerland and Arp 1993a) and B cells are not found in chicken lung until the second week after hatch (Jeurissen, Janse, Koch, et al. 1989). During the following weeks, increasing numbers of CD45+ cells accumulate and develop into organized structures with T cells in the centre and B cells in the periphery (Jeurissen et al. 1994). These organized BALT structures are not observed prior to the third to fourth week after hatching and GC are first seen in 3-week-old chickens (Fagerland and Arp 1990; Fagerland and Arp 1993a; Reese et al. 2006). Fully developed BALT, with B cells in the central area covered by a CD4+ cell cap, is found in birds that are 6 weeks of age and older (Fagerland and Arp 1990), and at that time mature antibody-secreting cells (ASC) of all isotypes are seen under or within BALT epithelium and in non-BALT areas (Jeurissen, JanseKoch, et al. 1989; Fagerland and Arp 1993a). By this time, nearly every bronchial opening is surrounded by the lymphoid tissue (Fagerland and Arp 1993b). As there is no local production of Ig in the chicken BALT until the first week after hatch and IgA is only produced after the second week, in young chickens Ig against respiratory pathogens must be serum derived. The Ig could be derived either from maternal sources, or be produced elsewhere (Fagerland and Arp 1993a).

Mature BALT consists of lymphocyte aggregates covered by a distinct layer of epithelium, harbouring considerable numbers of lymphocytes (Figure 3). This epithelium is referred to as follicle-associated epithelium (FAE). In mammalian mucosa-associated lymphoid tissues (MALT), the epithelial layer contains specialized M-cells, but these are not observed in chicken BALT (Fagerland and Arp 1993b). In FAE, cells are present that display some characteristics of M-cells, however particle uptake by these cells was not observed (Fagerland and Arp 1990; Fagerland and Arp 1993b). In 6–8-week-old birds, GC are found in most nodules and occasionally ASC are found under the FAE (Jeurissen, Janse, Kok, et al. 1989; Fagerland and Arp 1993a).

Figure 3. Section showing fully developed BALT. A: B cell staining. B: CD4 T cell staining (adapted from Kothlow and Kaspers 2008, with permission from Elsevier).

3.2. The phagocyte system of the RT

Following RT infection or vaccination, the antigen (Ag) needs to be taken up and presented to cells of the adaptive immune system. A variety of antigen-presenting cells (APC) is present in rodent and human RT including dendritic cells (DC), macrophages (MPh) and B cells. RT-DC are considered to be attractive targets for mucosal vaccination strategies, since a large network of phagocytes, including DC is located beneath the epithelium of several mucosal tissues, such as the airway and the intestines and they are potent activators of naive T cells.

In the mouse lung, DC are found in the large conducting airways, the lung parenchyma and in the alveolar space (de Heer et al. 2005); both conventional (c)DC and plasmacytoid (p)DC are present. cDC can be divided in trachea and large airway epithelial DC expressing CD11c, langerin and CD103, submucosal DC expressing CD11b and CD11c and interstitial DC that can be either CD11b+ or CD11b- (GeurtsvanKessel et al. 2008). In alveolar space, a population of alveolar MPh is also present. In the RT, tracheal DC have a rapid turnover rate (2–3 days), while interstitial DC are long lived (Holt et al. 1994; von Garnier et al. 2005).

The chicken RT contains few free-residing MPh (‘avian respiratory phagocytes’; ARP) compared to mammals and MPh are absent from the surface of the air capillaries (Maina 2002). In avian lung, MPh are present in the lining of the atria and infundibulae (Maina 2002) and are also abundantly present in the connective tissue (Klika et al. 1996). Phagocytic cells are strategically localized at the start of the gas-exchange area to clear the air of inhaled particles, before it reaches the thin and vulnerable air capillaries (Kothlow 2008). Most diffusely distributed leukocytes in interstitial lung tissue belong to the monocyte/MPh and DC phenotype, because of their expression of MHC-II, and their positive staining using the antibodies (Ab) CVI-ChNl-68.1 and −74.2 (Jeurissen, Jeurissen, Janse, Koch, et al. 1989; Jeurissen, Janse, Kok, et al. 1989). Adjacent to the muscle fibers that form the parabronchi, the lumen is bordered by 74.3 stained cells, and these cells are expected to act as mammalian alveolar MPh (Jeurissen et al. 1994). Epithelial cells lining the atrial muscles, the atrial floor and the infundibulae have been shown to be phagocytic and may therefore play a role in the defence of the thin, extensive and highly vulnerable tissue barrier (Nganpiep and Maina 2002). It is unclear if phagocytosed material is presented to the local immune system in the BALT, or in the follicles present in the interstitium between parabronchi or if it is transported to the spleen or a combination of these option depending on the antigen. In chicken, BALT is constitutively present and is thought to compensate for the lack of dLN. When chickens were inoculated with fluorescent beads that were coated with LPS as a bacterial Ag, bead + cells were present in BALT areas (de Geus, Jansen, et al. 2012). This could indicate that following the uptake of Ag, APC migrate mainly locally towards the BALT area. Moreover, bead + cells were rarely present in blood and spleen, suggesting that local induction of RT immune responses might dominate over the systemic induction.

Ag-bearing DC in the mammalian RT mainly migrate towards mediastinal lymph nodes (MLN), where they interact with naïve T cells (Legge and Braciale 2003; Hammad and Lambrecht 2007; GeurtsvanKessel et al. 2008; Bakocevic et al. 2010; Lambrecht and Hammad 2010). In chicken, it is not known which cell type is responsible for priming of naive T cells and whether a functional distinction between MPh and DC in priming capacity exists. In mouse, the endosomal compartment of DC has a near neutral pH (Savina et al. 2006; Savina et al. 2009), while MPh have much lower phagosomal pH (Yates and Russell 2005). In mouse, the decreased phagosomal acidification in DC reduces Ag proteolysis to a level that allows processing but does not fully destroy antigenic peptides. This contributes to the high Ag-presenting capacity of DC (Mantegazza et al. 2008; Ramachandra et al. 2009). MPh have much higher levels of proteolysis than DC, which limits their efficiency as APC (Delamarre et al. 2005). Recently, lung phagocytic cells were identified in chicken that decreased the pH of their endosomal compartment following the uptake of a bead (CD11 + , KUL01+ and CD40+ phagocytes) and cells that did not (MHC II+ and CD80+ phagocytes) (de Geus, Jansen, et al. 2012), suggesting that in chicken MPh also are functionally distinct from DC. Whether the capacity to present antigen differs between these cell populations needs further investigation. These findings can facilitate the specific targeting of DC using a mucosal vaccine.

3.3. Humoral immune responses in the RT

The mucosal tissue of the nose is first to come into contact with aerosols or micro-organisms during inhalation. Large particles and micro-organisms can be removed mechanically by the mucus, but invasive pathogens need to be controlled by the immune system. A major characteristic of organized chicken nasal-associated lymphoid tissue (NALT) is the formation of circumscribed B cell areas occasionally displaying GC, covered by a CD4+ T cell cap. In both organized NALT and in the epithelium, the dominant isotype is IgY, whereas IgM+ cells are less frequent and IgA+ cells are relatively rare (Ohshima and Hiramatsu 2000). IgY-containing cells are more frequently observed than the other isotypes, so in contrast to the importance of IgA in the mammalian mucosal immune system, IgY may be more important at avian mucosal surfaces, at least in the nasal cavity (Ohshima and Hiramatsu 2000).

The major eye-associated lymphoid tissues are located in the HG and in the conjunctiva of the lower eyelid. Secretions of the HG drain into the upper RT, providing local protection by secretory Ig (Dohms et al. 1981). Almost all tear Ig is produced locally in the HG, because removal of the HG abrogated IgM and IgA and reduced IgY in tears (Scott et al. 1993; Jeurissen et al. 1994). The HG is organized in two histologically-distinct compartments, a head and a body. The head of the gland resembles a secondary lymphoid organ, with lymphoid accumulation and GC. The majority of B cells are IgA+, although many IgY+ cells are found as well (Jeurissen et al. 1994). The plasma cell-filled body resembles a tertiary lymphoid structure (Olah et al. 1996). Discrepancies exist regarding Ig expression in HG. Olah et al. (1996) found that the ASC produce mainly IgA and IgM, whereas Jeurissen et al. (1994) showed the presence of a considerable number of IgY-producing cells. Locally produced Ig of all isotypes appears in tears together with IgY transudated from serum (Toro et al. 1993), thus protecting the upper RT. The ASC are present in comparable numbers in chickens raised under SPF and under conventional conditions (Bang and Bang 1968). In steady state situations, all Ig isotypes are produced, but after local immunization with tetanus toxoid vaccine, IgA and IgY were produced in high concentrations, with only trace amounts of IgM (Shirama et al. 1996). The ASC can migrate to the epithelium, where they make contact with cells resembling M cells. These cells are probably involved in Ig-secretion (Gallego and Glick 1988).

There is no constitutive lymphoid tissue present in trachea. However, the tracheal mucosa is highly responsive to infection and reacts with extensive lymphocyte infiltration followed by proliferation (Gaunson et al. 2000; Gaunson et al. 2006; Matthijs et al. 2009). Tracheal lesions characteristic for Mycoplasma infections predominantly consist of proliferating B cells (Gaunson et al. 2006) and this is similar to IBV infection (Kotani et al. 2000). The cells are mainly located in the lamina propria and submucosa (Kotani et al. 2000). Responses in trachea are age-dependent as B cells were not detected in the trachea of Mycoplasma gallisepticum-infected SPF layer-type chicken until 3 weeks post infection (Gaunson et al. 2006).

In interstitial lung tissue, IgM+ cells are found from 5 days after hatch (Jeurissen, Janse, Kok, et al. 1989), while IgA+ and IgY+ cells are not found at that time (Jeurissen, Janse, Kok, 1989b). In older birds, B cells are found throughout the parenchyma (Reese et al. 2006). In BALT and other regions of the lung, IgM-producing cells predominate over IgY-producing cells and IgA-producing cells (Fagerland and Arp 1993a). ASC are not present in the gas-exchange areas of the lung (Fagerland and Arp 1993a).

4. Induction of influenza-specific immunity

Induction of an antiviral adaptive immune response requires uptake of AIV by immature respiratory DC. In mice, both pDC and cDC are necessary for the induction of an adaptive response following influenza infection (GeurtsvanKessel et al. 2008). In mammals, Ag-bearing DC migrate to the MLN where they interact with T cells. It has been shown that following influenza infection, the langerin + CD103 + CD11b – cDC that migrated to the MLN presented Ag ex vivo to both CD4+ and CD8+ T cells (GeurtsvanKessel et al. 2008).

In AIV infections, humoral immune responses contribute to virus neutralization through binding to viral Ag. In humans, all currently used vaccines depend on the induction of neutralizing antibody responses (Letvin 2007). Humoral memory, i.e. secretion of specific neutralising antibodies over long periods in the apparent absence of the Ag provides the host with a first line of defence against reinfection (reviewed in Dorner and Radbruch 2007).

In influenza-infected mice, virus-specific ASC are detected in various organs, such as lung, MLN, bone marrow (BM) and spleen (Owens et al. 1981; Jones and Ada 1986; Joo et al. 2008; Wolf et al. 2011). In humans, HA-specific IgA in nasal washes is considered to be an important mediator in protection from influenza infection (Clements et al. 1986). Also in chicken, Ig are involved in protection against AIV as birds that are protected by vaccination have high serum titres of influenza-specific Ig (Kumar et al. 2007; Maas et al. 2009; Nayak et al. 2009). The role of respiratory Ig in AIV-specific immunity has previously been investigated after vaccination at the level of secreted Ig in tears and serum and ASC in HG. It is known that Ig are involved in protection against AIV as birds that are protected by vaccination have high serum titres of influenza-specific Ig (Kumar et al. 2007; Maas et al. 2009; Nayak et al. 2009). Van Ginkel et al. (2008) studied mucosal and systemic humoral immune responses following ocular influenza vaccination with a H5-expressing replication competent adenovirus (RCA)-free human E1/E3-defective Ad5 vector. High frequencies of H5-specific IgA ASC observed in HG- and H5-specific Ig were found in serum and tears. In influenza-infected chickens, IgM responses predominate during primary infection. Antiviral IgA and IgY responses were induced both locally in the lung and systemically in the spleen. Responses in the lung preceded responses in the spleen, indicating that respiratory immune responses were not only induced in the spleen but also locally in the lung. The AIV-specific ASC responses in lung significantly correlated to responses in bone marrow (BM), suggesting also a role for BM in respiratory immunity (de Geus, Rebel, et al. 2012).

In vivo, most ASC that are generated in a given immune reaction disappear from the secondary lymphoid organs within a few days to weeks. In a secondary immune response in mice, 10–20% of the ASC that are generated survive in the BM for more than 2 weeks. The persisting ASC can survive in vivo for long periods and can be detected in BM for at least the next 60 days (reviewed by Radbruch et al. 2006). Also in chickens, the BM could be a survival niche for long-living ASC. However, when chickens were infected with LP H7N1 AIV, serum responses and also AIV-specific ASC numbers in all organs, including BM, declined very rapidly and could not be detected at 3 weeks post primary and 6 weeks post secondary infection (de Geus, Rebel, et al. 2012). This might suggest that H7N1 infection generates short-lived ASC. Generation of short-lived ASC has been described in a mouse vesicular stomatitis virus model, where ASC were transferred into naïve mice. In the presence of Ag, high numbers of ASC are maintained; while in the absence of Ag, ASC numbers decline rapidly (Ochsenbein et al. 2000).

In mice, CD8+ cytotoxic T cells (CTL) also make an important contribution in influenza-specific immunity (Kim et al. 2011). In chickens, the role of influenza-specific CTL is less clear. Pulmonary CD8+ cells were shown to be important in protection against AIV (Seo et al. 2002) and cells isolated from lungs of AIV-infected chickens secrete IFN-γ after ex vivo restimulation (Rauw et al. 2011). Furthermore, in a recent study, MHC I-inbred chickens were infected with AIV and cells isolated from lung were stimulated ex vivo with several predicted CD8 + T cell epitopes. These epitopes where predicted, based on anchor residues described for the different MHC I types of the MHC I-inbred chickens. This way, novel influenza-specific CD8 + T cell epitopes were identified in MHC I-inbred chickens (Reemers et al. 2012).

In conclusion, both humoral and cellular responses play a role in protection from influenza infection. Neutralizing Ab are crucial, as chickens with measurable HI titres after vaccination were protected from clinical disease following AIV infection (Kumar et al. 2007; Maas et al. 2009). Furthermore, passive immunization with AIV protein-specific antiserum protected birds from challenge infection (Shahzad et al. 2008).

5. Vaccination strategies

The risk and the size of an outbreak of AIV could be restricted by vaccination of poultry. Currently licensed vaccines for use in chicken are adjuvanted whole-inactivated virus (WIV) or split virus, inactivated reverse-genetics virus for H5N1 and H5N3, and fowl pox and NDV vector vaccines (reviewed by van den Berg et al. 2008). Possible vaccination strategies are rapid intervention (e.g. emergency vaccination of poultry in the area around an outbreak), preventive vaccination of specific categories of poultry that are more at risk for a new introduction of AIV (e.g. free range layers in areas with many wild ducks and geese) and general preventive vaccinations of poultry in area's in which AIV is endemic. A vaccine used for rapid intervention during an AIV outbreak should be safe, highly effective after a single administration and be suitable for mass application. A vaccine that could be applied by spray or aerosol would be suitable for mass application, which is regularly performed, e.g. IBV (De Wit et al. 2010), NDV, avian metapneumovirus and most Mycoplasma galliseptum vaccines (Ley 2003). In the case of AIV, aerosol vaccination using live virus is not desirable because of its zoonotic potential and because of the risk for virus reassortment. Mucosal vaccination via the respiratory route has several advantages: it induces both local and systemic immune responses (Atmar et al. 2007; Tseng et al. 2009; Worrall et al. 2009), it could halt infection already at portal of entry (Yoshikawa et al. 2004) and it is suitable for mass application.

5.1. Mucosal application of inactivated vaccines

Intranasally (i.n.) applied whole inactivated AIV is poorly immunogenic (Hagenaars et al. 2008; Worrall et al. 2009; de Geus et al. 2011) and this is also described in chicken for i.n. applied inactivated NDV (Tseng et al. 2009). To enhance the immunogenicity of whole-inactivated AIV it needs to be adjuvanted. In the field of mucosal vaccination advances have been made in the development of mucosal adjuvants that protect the Ag from degradation and can increase nasal residence time, such as poly(lactic-co-glycolic acid) (PLGA) and chitosan (and its derivative trimethyl chitosan; TMC) (Slutter et al. 2010; Keijzer et al. 2011).

The local induction of RT tract immune responses in chicken makes it attractive to apply a vaccine to the RT. However, when different adjuvants were tested in an aerosolized vaccine using WIV, none of the adjuvanted vaccines induced AIV-specific Ig after single vaccination, measured 1 and 3 weeks after vaccination by aerosol, in contrast to the intramuscularly applied vaccine. The aerosolized vaccine did enter the chickens’ RT as cholera toxin B subunit (CT-B)-specific serum Ig were detected after 1 week in chickens vaccinated with the CT-B-adjuvanted vaccine (de Geus et al. 2011). Also using an i.n. vaccination strategy, no AIV-specific Ig was found until 3 weeks after the third vaccination (de Geus 2012). Although the respiratory route is ideal for mass application, for the use of whole inactivated virus further modifications of a vaccine are needed, for example with regard to retention time and to efficiently induce an immune response in the tolerizing environment of the RT. Van Eck (1990) vaccinated SPF chickens repeatedly with inactivated NDV in PD, a carbomer adjuvant. Only low HI titers were induced after oculonasal vaccination and several booster vaccinations did not result in increased titers. Furthermore, very high doses of antigen were required to induce protection from challenge infection. Following i.n. application, approximately 70–80% of the applied volume is swallowed and thus ends up in the gastrointestinal tract (Van Eck 2011, personal communication). This could affect immunogenicity of WIV, while leaving immunogenicity of CT-B intact. As no AIV-specific Ig was induced both after aerosol vaccination and after repeated i.n. vaccination, future research for rapid intervention strategies or mucosal vaccination strategies should focus on alternatives such as the use of live vector or subunit vaccines. Other approaches could include enhancing retention time of the WIV vaccine and increasing Ag uptake by targeting RT-DC subsets.

5.2. Alternative vaccination strategies suitable for mucosal application

Adjuvanted inactivated AIV did not induce detectable levels of serum Ab after both aerosol and i.n. applications (de Geus et al. 2011). An alternative approach for mass vaccination of chickens could be the use of live vector vaccines. Vector vaccines can be compatible with Differentiating Infected from Vaccinated Animals (DIVA) strategies. The currently licensed vector vaccines for influenza vaccination in chicken are recombinant fowl pox virus and NDV vector vaccine. The recombinant fowl pox vaccine induced good protection from heterologous challenge (Swayne et al. 2007), but can interfere with preexisting immunity against fowl pox. HA-expressing NDV vector vaccines can be applied by spray or in drinking water and have shown to protect against challenge infection in chicken (Swayne et al. 2003; Nayak et al. 2009; Schroer et al. 2011). Similar to the fowl pox vaccine, the NDV vector vaccine can also interfere with preexisting NDV immunity induced by NDV vaccination. Furthermore, the presence of maternal Ab can interfere with vaccine efficacy. Other vectors that are currently being tested for mucosal vaccination strategies include adenovirus Ad5, infectious laryngotracheitis (ILT) recombinants and baculovirus. Mucosally applied AIV protein-expressing Ad5 induces high levels of AIV-specific Ig and also HI titers against the whole influenza virus in mouse (Price et al. 2010), ferrets (Roy et al. 2011) and chicken (van Ginkel et al. 2008). Coarse spray vaccination of chickens did not induce any adverse effects and slightly elevated H5-specific IgA was found in tears (Toro et al. 2010). Ocular (Veits et al. 2003; Pavlova et al. 2009) or tracheal (Luschow et al. 2001) vaccination with HA-expressing ILT induces HA-specific Ig and protects from challenge infection. Vaccination with H7-expressing baculovirus induced H5-specific Ig after intramuscular vaccination, but responses after i.n. vaccination were much lower (He et al. 2008).

In summary, a vector vaccine has to be chosen with care because interference with existing vaccination programs or maternal antibodies against the viral vectors (in the case of, for instance, NDV and fowl pox) could interfere with the induction of protective Ig responses. Furthermore, the choice of the vector is important because a viral vector might reassort and could pose a health threat. Although rarely a problem, at least for the NDV vector the possibility of occurrence of reassortment is under debate (Collins et al. 2008; Han et al. 2008).

Challenges in respiratory vaccination include the mucociliary clearance mechanisms in the nasal cavity and the trachea, resulting in a low residence time and the low permeability of the mucosa (Ugwoke et al. 2001; Illum 2003). Mucosal residence time can be improved by using particles with mucoadhesive properties (Smart 2005; Jaganathan and Vyas 2006). These particles are generally inert, but provide persistent Ag exposure and could facilitate the uptake by APC (reviewed by O'Hagan and Valiante 2003). Further advantages of mucoadhesive particles include the possibility to crosslink the Ag on the particle surface and the possibility to include immunomodulatory agents such as TLR ligands, thereby increasing recognition and immunogenicity (reviewed by O'Hagan and Valiante 2003; Sharma et al. 2009). Enhancing the residence time in the RT of a vaccine would prolong its exposure to RT-DC, resulting in increased uptake and Ag-presentation. As respiratory administration of inactivated AIV alone does not result in upregulation of MHC II and CD80 expression by RT-DC (de Geus, Jansen, et al. 2012), it will be necessary to co-adjuvant with or co-administer a mucoadhesive particle. Particles that are currently tested in mucosal vaccines include PLGA spheres or chitosan and its derivative TMC.

PLGA spheres consist of biocompatible copolymers and are approved for use in humans. A major problem in the use of PLGA spheres is the protein instability of incorporated Ag (van de Weert et al. 2000). This problem could be solved by adsorbing Ag onto the surface of the particle (Kazzaz et al. 2000). In mice, PLGA particles are poorly taken up by DC and do not induce subsequent maturation. Furthermore, nasal administration of OVA-PLGA spheres did not induce specific Ig responses, even after tertiary vaccination (Slutter et al. 2010). In turkeys, PLGA particles are taken up by MPh in vitro, but oculonasal vaccination with live avian metapneumovirus (aMPV) was far superior to vaccination with aMPV F protein-loaded PLGA as live aMPV vaccination completely protected birds from clinical disease and only 50% of aMPV F protein-PLGA vaccinated birds were protected from challenge infection (Liman et al. 2007).

TMC is a chitosan derivative that is more stable at neutral pH than chitosan itself and has shown promising results in i.n. vaccines in mice (Hagenaars et al. 2009; Slutter et al. 2010). Chitosan and TMC have mucoadhesive properties and have the ability to cross transepithelial barriers. Besides its mucoadhesive properties, chitosan can also act as an adjuvant (Ghendon et al. 2008; Ghendon et al. 2009). TMC significantly increases nasal residence time of co-administered OVA and high levels of IgG and sIgA were induced after nasal administration (Slutter et al. 2010). Both adjuvants should therefore be able to overcome the challenges posed by nasal administration. Chitosan has shown promising results in i.n. split virus vaccines in mice (Bacon et al. 2000), split virus and protein vaccines in humans (Read et al. 2005; Sui, Chen, Fang, et al. 2010; Sui, Chen, Wu, et al. 2010) and split virus and DNA vaccines in poultry (Tischer et al. 2002; Worrall et al. 2009; Huang et al. 2010; Rauw et al. 2010). However, in our study vaccination with chitosan-adjuvanted inactivated AIV and TMC inactivated AIV particles did not induce AIV-specific serum Ig (de Geus et al. 2011). It was published that chitosan can work synergistically with secondary adjuvants such as CT-B (Moschos et al. 2004). It would therefore be worthwhile to co-adjuvant a chitosan- or TMC-based AIV vaccine for use in chicken.

In summary, mucoadhesive particles are promising vaccine components for use in an AIV vaccine for rapid intervention strategy, but studies are needed to properly co-adjuvant these particles and to prevent Ag degradation.

6. Concluding remarks

Respiratory immune responses in the chicken are locally induced. This would be advantageous for mucosal vaccination strategies. Possible mucosal vaccination strategies for general preventive vaccination include the use of adjuvanted inactivated AIV and the use of vector viruses. Furthermore, vaccination strategies could include the use of particles that increase mucosal residence time or the specific targeting of vaccine Ag to RT-DC subsets. Targeting or enhanced recruitment of RT-DC subsets could be achieved by adding a specific adjuvant, making use of particulate Ag-delivery systems that promote phagocytosis or by the targeting of receptors expressed by RT-DC, such as glycan-recognizing receptors. For respiratory application, particles should be designed in such a way that they can also reach the lung tissue itself to be taken up by RT-DC. Particles in size ranging from 1 to 3.7 µm can be expected to distribute throughout the RT (Hayter and Besch 1974; Hayter 1977; Corbanie et al. 2006; Tell et al. 2006). When a vaccine is designed for a rapid intervention strategy, a possible approach could be the activation of the innate immune system by, for instance, CT-B (Matsuo et al. 2000), resulting in a partial protection until the adaptive immune system is activated.