Exploring the host transcriptome for mechanisms underlying protective immunity and resistance to nematode infections in ruminants

Veterinary Parasitology 190 (2012) 1–11

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Veterinary Parasitology journal homepage: www.elsevier.com/locate/vetpar

Review

Exploring the host transcriptome for mechanisms underlying protective immunity and resistance to nematode infections in ruminants Robert W. Li a,∗ , Ratan K. Choudhary a , Anthony V. Capuco a , Joseph F. Urban Jr. b a b

USDA-ARS, Bovine Functional Genomics Laboratory, Beltsville, MD, USA USDA-ARS, Diet, Genomics, and Immunology Laboratory, Beltsville, MD, USA

a r t i c l e

i n f o

Article history: Received 19 March 2012 Received in revised form 8 June 2012 Accepted 15 June 2012 Keywords: Ruminant Nematode Resistance Parasite Pathways Transcriptome

a b s t r a c t Nematode infections in ruminants are a major impediment to the profitable production of meat and dairy products, especially for small farms. Gastrointestinal parasitism not only negatively impacts weight gain and milk yield, but is also a major cause of mortality in small ruminants. The current parasite control strategy involves heavy use of anthelmintics that has resulted in the emergence of drug-resistant parasite strains. This, in addition to increasing consumer demand for animal products that are free of drug residues has stimulated development of alternative strategies, including selective breeding of parasite resistant ruminants. The development of protective immunity and manifestations of resistance to nematode infections relies upon the precise expression of the host genome that is often confounded by mechanisms simultaneously required to control multiple nematode species as well as ecto- and protozoan parasites, and microbial and viral pathogens. Understanding the molecular mechanisms underlying these processes represents a key step toward development of effective new parasite control strategies. Recent progress in characterizing the transcriptome of both hosts and parasites, utilizing high-throughput microarrays and RNAseq technology, has led to the recognition of unique interactions and the identification of genes and biological pathways involved in the response to parasitism. Innovative use of the knowledge gained by these technologies should provide a basis for enhancing innate immunity while limiting the polarization of acquired immunity can negatively affect optimal responses to co-infection. Strategies for parasite control that use diet and vaccine/adjuvant combination could be evaluated by monitoring the host transcriptome for induction of appropriate mechanisms for imparting parasite resistance. Knowledge of different mechanisms of host immunity and the critical regulation of parasite development, physiology, and virulence can also selectively identify targets for parasite control. Comparative transcriptome analysis, in concert with genome-wide association (GWS) studies to identify quantitative trait loci (QTLs) affecting host resistance, represents a promising molecular technology to evaluate integrated control strategies that involve breed and environmental factors that contribute to parasite resistance and improved performance. Tailoring these factors to control parasitism without severely affecting production qualities, management efficiencies, and responses to pathogenic co-infection will remain a challenge. This review summarizes recent progress and limitations of understanding regulatory genetic networks and biological pathways that affect host resistance and susceptibility to nematode infection in ruminants. Published by Elsevier B.V.

∗ Corresponding author at: Animal and Natural Resources Institute, USDA-ARS, 10300 Baltimore Avenue, Beltsville, MD 20705, USA. Tel.: +1 301 504 5185; fax: +1 301 504 8414. E-mail address: [email protected] (R.W. Li). 0304-4017/$ – see front matter. Published by Elsevier B.V. http://dx.doi.org/10.1016/j.vetpar.2012.06.021

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Contents 1. 2. 3. 4. 5. 6.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cytokines and cell adhesion molecules in host immune responses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Transcriptome characteristics of the bovine GI tract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Biological pathways affected by nematode infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Bridging QTL and transcriptome analyses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction Nematode infections in ruminants have important economic consequences. The most economically relevant parasitic nematodes that infect the gastrointestinal (GI) tracts of ruminants include species from Cooperia, Haemonchus, Nematodirus, Oesophagostomum, Ostertagia, Trichuris, and Trichostrongylus. Economic losses from nematode infections in ruminants include reduction in milk and meat production and increases in production costs that include prophylactic measures for reducing infection, treatment of infected animals, and mortality. The precise monetary loss is difficult to determine (Charlier et al., 2009); however it is estimated that the annual loss to the U.S. cattle industry due to parasitic infections of the GI tract is close to $2 billion. The pathogenic effects of infection that cause economic loss are often associated with the severity of the disease. A mortality of as high as 20% of young goats is attributed to GI parasites (Valentine et al., 2007). However, the most common symptoms of parasitic infection of the GI tract are compromised production traits, including reduced weight gains (Coop et al., 1979) and milk yield (up to 2.2 kg of milk per animal per day) in cattle (Barger and Gibbs, 1981). Ostertagia ostertagi is arguably the most important cattle parasite in temperate regions of the world (Li et al., 2010). In Canada, France, and Belgium an increase in O. ostertagi specific antibody in milk is associated with a decrease in the annual average milk production by 1.2, 0.9 and 1.2 kg/cow/day, respectively (Charlier et al., 2005; Sanchez and Dohoo, 2002). Conversely, anthelmintic treatment is often associated with a positive milk response (Sanchez et al., 2004). In addition, nematode eggs shed in the feces of the host (fecal egg counts or FEC) are an indicator of the level of infection and are negatively correlated to the live body weight of Tswana goats (Nsoso et al., 2001). The parasitic nematodes species from the genera Haemonchus, Ostertagia, Trichostrongylus, Nematodirus, and Cooperia are ubiquitous and are often associated with production losses and clinical symptoms. One report from Northeastern China indicates that administration of de-wormers tends to increase the average body weight (2.6–10.8%) and wool production (1.7%), as well as to decrease death rate up to 10% in sheep (FAO, 2002). The mainstay of parasite control for the past five decades has been use of anthelminthics. However, indiscriminate use of anthelmintics has led to widespread parasite drug resistance and resulted in a shift in the most prevalent populations of parasitic nematodes on pasture (Gasbarre et al., 2009). Resistance to anthelmintics of the benzimidazole and macrocyclic lactone class has been

2 3 5 6 8 8 9 9

reported around the world (Hughes et al., 2005). The issue of drug resistance has become a global concern (Mortensen et al., 2003). High prevalence of drug-resistant parasites in ruminants has called for an urgent re-examination of our current treatment strategies and for the development of alternative means to control parasitic nematodes that limit ruminant productivity. Three general approaches are available for controlling infection by disrupting the life cycle of the nematode and reducing the severity of infection (Hoste and TorresAcosta, 2011). These include: (1) grazing management to reduce the number of infective nematodes and host exposure, (2) elimination of nematodes in the host by conventional anthelminthics and alternative diets enriched in phytochemicals, and (3) enhancing host resistance by immune therapies or selective breeding. This review focuses primarily on the latter approach with an emphasis on the use of transcriptome analyses to provide potential targets for immune therapies and selective breeding. Although predictions for effective vaccines against ruminant GI nematodes have not been fully realized (Emery et al., 1993), optimism has been renewed by the application of gene discovery to identify parasite antigens that can induce host protective responses. For example, vaccines against Haemonchus contortus in sheep (LeJambre et al., 2008), goats (Ruiz et al., 2004) and cattle (Bassetto et al., 2011) have been developed but have had limited success because of cost and ineffective application. The dose, frequency, and timing of vaccine delivery, and appropriate adjutants required to enhance efficacy need to be balanced against reduced profitability. Furthermore, vaccines that confer protection against all major parasite species in a given locality are still not on the horizon. Defined mechanisms of resistance that are amenable to active immune therapies are needed to provide an appropriate protective response to nematodes without negative consequences to the control of by-stander pathogens (Finkelman and Urban, 2001). Coupled with disease reduction and enhanced host resistance to GI nematodes is the nutritional status of the animal. Infection with GI nematodes is accompanied by mild to severe hypophagia, impaired digestion and absorption, and increased partitioning of nutrients toward immune function (Hoste and Torres-Acosta, 2011; Sykes, 2008). Reduced appetite during GI parasite infection of lambs was shown to be mediated by cholecystokinin (Dynes et al., 1998). In mice, appetite depression was immunologically linked to secretion of cholecystokinin by enteroendocrine cells of the gut mucosa mediated

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by Th2 cytokines IL4 and IL13 (McDermott et al., 2006). Nutrient requirements for these infections are related to the cost of repairing damaged gut mucosa and mounting an immune response. The latter cost predominates, due to the extensive replication of lymphocytes and other immune cells, as well as the synthesis of acute phase proteins (Colditz, 2008). It is logical that nutrient management can improve clinical symptoms and enhance the host’s ability to resist infection by GI nematodes (Wagland et al., 1984). Because the impact of infection on protein metabolism typically exceeds its impact on energy balance, the resilience and productivity of infected animals can be improved by protein supplementation (Coop and Kyriazakis, 1999). Furthermore, studies in rats and sheep showed supplemental dietary protein induced transcriptome changes that increased cell turnover, cell growth and differentiation, and expulsion of nematodes from the intestine (Athanasiadou et al., 2011; Keane et al., 2006). Elucidation of the metabolic costs of parasitism and appropriate nutritional supplementation to improve the health and performance of infected and noninfected animals remains an important need. Coupling dietary changes to transcriptomic expression of appropriate resistance mechanism could improve the efficiency of integrated strategies for parasite control. Enhanced resistance to nematode infections that improves ruminant health and vigor includes reduced penetration of parasitic larvae and persistence of developmentally arrested larvae, decreased worm fecundity, accelerated worm expulsion, and reduced pathology. Considerable evidence supports the idea that traits associated with parasite resistance are under host genetic control (Gasbarre et al., 1993, 2001). Resistance to infection likely results from inheritance of genes or genomic loci that have a direct or indirect role in expression of molecules that appropriately regulate host immunity to control infection and limit pathology. Variants in these genes and/or genomic loci can be applied to breeding programs that enhance parasite resistance. Heritability is trait-dependent. For example, heritability of adult worm length at the end of first grazing season is high (h2 = 0.62) in sheep (Stear et al., 2009) while heritability for FECs is moderate in sheep (Albers et al., 1987), goats (Mandonnet et al., 2001), and cattle (Gasbarre et al., 2001). Anticipated advantages of selective breeding, utilizing genetic markers associated with resistance traits, have spurred research efforts (Amarante et al., 1999; Andronicos et al., 2010; Burke and Miller, 2004; Crawford et al., 2006) to decrease anthelmintic residues in animal products and the environment as well as increasing profitability of animal production. As a result, several programs, such as Sustainable Control of Parasites in Sheep (SCOPS) and Control of Worms Sustainably (COWS) have been recently initiated (Taylor, 2012). Understanding the molecular mechanisms that contribute to protective immunity, immune suppression, pathology, and host resistance will have a huge impact on alternative control strategies. In this review, we discuss outcomes of the recent application of high-throughput genomic tools to further our understanding of regulatory genetic networks and biological pathways that define host

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resistance and susceptibility to GI nematode infection in ruminants.

2. Cytokines and cell adhesion molecules in host immune responses Parasitism inherently reveals strategies employed by the parasite to survive in the host. Parasites have the ability to modulate host immune responses and sustain long-lasting chronic infection by escaping host immune surveillance and modulating host protective mechanisms. The hallmark of nematode infections is T-helper (Th0) activation in naïve animals that develops into a robust Th2 response characterized by a particular pattern of cytokines and related cell activation pathways. Defining cytokine responses to infection may contribute to the development of successful vaccines and provide insight into protective immunity. In addition, a better understanding of the protective role of Th2 cytokines in nematode infections could improve efficacy through immune intervention. A caveat is that nematodes can also induce various levels of a Th1 response, but strongly polarized Th2 responses can down-modulate Th1-based immunity that is important for the control of protozoan and microbial co-infections (Finkelman and Urban, 2001). Studies using murine models demonstrate that GI nematode parasites induce CD4+ T cell responses skewed toward development of Th2 cells. These cells produce IL-4, IL-5, IL-9, IL-13, IL-25, and IL-33 which, in concert with stem cell factor and TFG-␤1, cause differentiation and maturation of intraepithelial mast cells, eosinophilia, and goblet cell development (Artis and Grencis, 2008). These events lead to alteration of enterocyte permeability, increased enterocyte turnover and the activation of goblet cells. Activation of goblet cells by IL-13 increases secretion of mucus and prevents contact of parasites with the epithelial surface, whereas other goblet cell products like Muc5A and Relm-beta are associated with anti-parasitic responses (Artis et al., 2004; Hasnain et al., 2011) and contribute to local inflammation in the mucosa (Li et al., 2007; Nair et al., 2008). Furthermore, IL-13 in conjunction with IL-4 activates macrophages that entrap and metabolically stress larval stage nematodes within the intestinal submucosa (Anthony et al., 2006; Artis and Grencis, 2008). Activated macrophages have also been shown to alter intestinal smooth muscle contractility and epithelial cell function in mouse models (Zhao et al., 2008). An accelerated turnover of enterocytes at the site of nematode infection has been identified as a mechanism of parasitic expulsion from the host, a mechanism that is regulated by local cytokines (Cliffe et al., 2005; Zaiss et al., 2006). However, even the strongly polarized intestinal Th2 response induced by infection of mice with Nippostrongylus brasiliensis express a regionally mixed immune response represented by brief Th17-induced neutrophil component in the lungs during larval migration (Chen et al., 2012). The expression of a Th1response during nematode infection can also determine the relative resistance versus susceptibility of a genetically distinct mouse strain (Else et al., 1992), and different nematode strains can induce different host cytokine patterns

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that contribute to the level of parasite burden (Johnston et al., 2005). In ruminants, a Th2-type of immune response induces high-level production of IgA and IgG, tissue eosinophilia, mucosal mast cell activation, and goblet cell hyperplasia to GI nematode infection (Li and Gasbarre, 2009; Li et al., 2010). For example, eosinophils generally comprise <5% of total leukocytes (Holtenius et al., 2004) but increase rapidly at the site of infection as the infection progresses (Li and Gasbarre, 2009). Haemonchus placei elicits a classical skewed Th2-type immune response in cattle. Gene expression of IL-4 and IL-13 is strongly up-regulated in resistant groups of Nellore cattle in comparison to susceptible groups (Zaros et al., 2010). In a field study, resistant Nellore cattle showed high serum levels of IgE induced by natural H. placei and C. punctata infections (Bricarello et al., 2007). However, infection of cattle with O. ostertagi elicits an immune response with features of concurrent Th1- and Th2-type responses. These infections induced a significant increase in IL-10 and IL-4 in abomasal lymph nodes (ABLN), the primary site of antigen presentation, by 11 days post infection (dpi) along with increased expression of IFN-␥ transcripts, which were up-regulated ∼25-fold over uninfected cattle and remained elevated through 28 dpi (Almería et al., 1997; Canals et al., 1997; Gasbarre et al., 2001). Cytokine gene expression by lymphocytes in the GI lamina propria of cattle after primary infection follows a similar pattern to the response in the ABLN: a strong upregulation of both IL-4 and IFN-␥ in the infected cattle at 10 dpi as well as in an established infection at 60 dpi. ABLNs undergo a drastic shift in the population profiles of lymphocytes and a rapid expansion of certain cell types after O. ostertagi infections. An increase in both IgM+ (B cells) and TcR1+ cells, as well as a reduction in the percentage of T cells, was observed after infection. The increase in B cell populations is consistent with the strong serum antibody responses and the increased IL-4 production that is elicited by parasite infection (Canals et al., 1997); and the increase in TcR1+ cell populations may be responsible for the observed up-regulation of IFN-␥ mRNA. After weeks of trickle infections with Ostertagia, the ABLNs from bull calves displayed attributes of a conversion from a Th1 dominant response to a Th2 dominant response, characterized by a reduction in the expression of IL-12 and IFN-␥, and a strong increase in IL-4, IL-5, IL-10, and IL-13 (Claerebout et al., 2005). A similar response to trickle infection has been observed in mice with a normally chronic infection with Heligmosomoides bakeri (Brailsford and Behnke, 1992). However, the abomasal mucosa of these infected cattle maintained attributes of concurrent Th1 and Th2 responses, characterized by strongly increased expression of IL-4, IL-10, and IFN-␥. These studies suggest that features of a Th1 response are exhibited for an extended period such that the Th1-type response is concurrent with the Th2 response during an established infection. While significant progress has been made in understanding cytokine responses during nematode infection in ruminants, many questions remain unanswered. Biological functions of cytokines contributing toward the development of protective immunity and host resistance

remains largely speculative. A vigorous and effective mucosal immunity is essential for the development of resistance to nematode infection in ruminants. Indeed, resistant sheep are able to more rapidly up-regulate Th2 cytokines than susceptible sheep (Terefe et al., 2007). However, cytokine expression may not exclusively correlate with resistance (Maizels and Yazdanbakhsh, 2003). For example, in Ostertagia-infected cattle no significant correlation has been found between levels of Th2 cytokines and common effector mechanisms in the mucosa, such as parasite-specific IgA, IgG, and IgM and the number of mast calls or eosinophils (Claerebout et al., 2005). In addition to cytokines, cell adhesion molecules appear to play an important role in eliciting a host response to GI nematodes. Cell adhesion molecules are essential for mediating cell–cell and cell–extracellular matrix interactions. These molecules, such as cadherins, integrins, lectins, and neural cell adhesion proteins, are involved in numerous biological processes, including cell proliferation and differentiation, tissue construction and wound repair, pathogen recognition and host defense (Li and Gasbarre, 2010). The recruitment of inflammatory cells to the site of infection is crucial for mounting a rapid and effective immune response, and can also affect the expression level of the Th1 response to a nematode infection (Bell and Else, 2008). Integrins play an important role in this process. For example, integrin-␤7 is associated with homing of mast cells to the mucosa of the small intestine (Pennock and Grencis, 2006). Lectins are important activators of the immune response and are associated with inflammation (Lasky, 1991). Galectins, intelectins and C-type lectins recognize surface carbohydrates present on parasites. Galectins have been suggested to play an important role in innate immunity, including serving as receptors for pathogen-associated molecular patterns (PAMP), activating various immune cells, and participating in cytotoxicity. Induction of galectin-11 expression in the GI tract of cattle infected by protozoa and nematodes has been reported (Hoorens et al., 2011). In addition, galectin-11 is strongly induced in the bovine GI tract by various parasites, such as O. ostertagia, Cooperia oncophora, and the protozoan parasite Giardia duodenalis (Hoorens et al., 2011), and in sheep by infections with H. contortus and Trichostrongylus vitrinus (Dunphy et al., 2000). Intelectin 2 (ITLN2) expression is regulated by IL-4 (French et al., 2007). Its elevated expression is observed in the sheep abomasum in response to Teladorsagia circumcincta infection, Dictyocaulus filaria natural infection (French et al., 2009), and H. contortus infection (Rowe et al., 2009). Most pertinent, this gene is naturally deleted in the genome of the susceptible mouse strain, C57BL/10, but is present in the genome of a nematoderesistant mouse strain, BALB/c, suggesting that ITLN2 may serve a protective role in the innate immune response to parasite infection (Pemberton et al., 2004). Indeed, the relative abundance of ITLN2 is significantly higher in the abomasum of resistant heifers than in that of susceptible heifers in response to Ostertagia infection (Li et al., 2011b), and in the abomasal mucosa of immune vs. naïve sheep in response to challenge with T. circumcincta (Athanasiadou et al., 2008). Cattle infected with Cooperia show strong up-regulation of additional cell adhesion molecules like

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cadherin 26 (CDH26), collectin-43, collectin-46, integrin, alpha 4 (ITGA4), and lectin, galactoside-binding, soluble, 13 (LGALS13) in the small intestine (Li and Gasbarre, 2009). Moreover, high expression of CDH26 is strongly correlated with eosinophils and FEC. Of note, CDH26 is a member of cadherin superfamily, which mediates cell–cell interactions in a calcium-dependent fashion. Cattle-specific collectins have been suggested to provide the first line of defense against pathogens without eliciting a general inflammatory reaction (Hansen et al., 2002). Collectins, upon recognition of PAMPs, cause opsonization, neutralization, agglutination, and phagocytosis of pathogens (Gupta and Surolia, 2007). Together, evidence suggests that cell adhesion molecules are involved in recognition of cell surface carbohydrate moieties of nematodes and play an important role in evoking effective host resistance to nematode infection. 3. Transcriptome characteristics of the bovine GI tract The abomasum and small intestine are the major predilection sites of parasitic nematodes in ruminants, including species from several genera such as Haemonchus and Ostertagia (the abomasum) and Cooperia (the small intestine). Understanding characteristics of the host transcriptome should facilitate identification of molecular mechanisms that underlie host resistance and protective immunity, especially when comparing animals with different resistant phenotypes. In addition, quantification of the transcriptome of both host tissues and parasites using nextgeneration sequencing technologies and bioinformatics has been initiated with the expectation that this provide insights into specific interactions between the host and pathogen genomes (Li et al., 2011a; Li and Schroeder, 2012). Recently, we have generated approximately 24 million 36-bp RNA-sequences per sample for abomasal and small intestine samples of 1-year-old calves (N = 6 and 8 for the abomasum and small intestine, respectively; unpublished data). These sequences were first checked for quality, and low-score nucleotides were trimmed using SolexaQA (Cox et al., 2010). The resultant sequences were analyzed using TopHat (Trapnell et al., 2009) and Cufflink (Trapnell et al., 2010). The number of genes expressed (supported by at least 1 sequence) was 18,930 ± 524 (mean ± sd) and 19,180 ± 258 in the abomasum and small intestine, respectively. The ENSEMBL bovine genebuild release 65 was used to identify 21,435 of the 24,616 genes (protein-coding as well as various RNA genes) collectively transcribed in the abomasum and small intestine. Our results indicated that the bovine abomasum and small intestine displayed distinct transcriptome characteristics. In the abomasum, the transcriptome was dominated by a handful of transcripts. An unknown gene (ENSBTAG00000047264) was the most abundant gene, accounting for 42.12% of all sequence reads, followed by centrosomal protein of 120 kDa (4.44%), ADAMTS-like 1 (4.40%), a novel gene (ENSBTAG00000031575, 3.65%), and aniline, actin binding protein (2.10%). Consistent with the previous study (Li et al., 2011b), immunoglobulin lambda-like polypeptide 1, polymeric immunoglobulin receptor (PIGR),

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complement C3, fatty acid-binding protein, liver, and elongation factor-1-alpha-1 were among the most abundant. The 50 most abundant genes accounted for 69.07% of all sequences from the abomasal transcriptome. In contrast, the 50 most abundant genes in the small intestine only accounted for 11.33% of all sequences. Of the 50 most abundant genes, 16 were shared by the transcriptomes of both tissues, including apolipoproteins A-I, A-IV, and B, CD74 antigen, complement C3, cytochrome c oxidase subunit 1, deleted in malignant brain tumors 1 (ENSBTAG00000022715, the most abundant gene in the small intestine, 2.26%), immunoglobulin J chain, and PIGR. Identification of tissue-specific expression signatures has both theoretical and pragmatic implications toward understanding host–parasite interactions. Whole transcriptome comparison between the abomasum versus small intestine identified five protein-coding genes that were uniquely expressed in one of the tissues. As shown in Table 1, ␣-(1,3)-fucosyltransferase (FUT9) and 2 unknown genes were transcribed in each abomasal sample but not detectable in the small intestine at the sequence depth used in this study, whereas two genes, including olfactory receptor, family 2, subfamily AE, member 1 (OR2AE1), were only expressed in the small intestine. FUT9 has much stronger activity for Lewis antigen biosynthesis than other members of ␣-(1,3)-fucosyltransferases and guides the expression of the CD15 epitope in epithelial cells of the GI tract (Nakayama et al., 2001). Interestingly, Helicobacter pylori, a Gram-negative bacterium colonizing gastric mucosa, also expresses Lewis antigen in the O chains of its LPS (Moran, 2008). The structural mimicry between the host and pathogen Lewis antigens may play an important role in gastric adaptation of H. pylori and modulation of the host innate immune response. It is foreseeable that unique expression of FUT9 in the bovine abomasum may have implications in host–pathogen relations. For example, uniquely expressed genes in a tissue could supply the unique attributes of a tissue niche that facilitate parasite colonization. Interference with parasite recognition of these cues could thus provide a basis for strategies to control parasitism that could preclude inflammation and immune modulating cascades induced by developing parasitic nematodes. The relative abundance of 1064 transcripts (including 854 protein-coding genes) differed between the abomasum and small intestine at a stringent false discovery rate <5%. For example, among the protein-coding genes, the abundance of mRNA for gastric inhibitory polypeptide (GIP) was 3-fold higher in the small intestine than the abomasum, consistent with previously published results (Musson et al., 2011). On the other hand, at least five lectins were more highly expressed in the abomasum than in the small intestine. For example, ITLN2 was ∼262-fold higher in the abomasum and collectin-43 and -46 (CL43 and CL46) were 268- and 157-fold higher, respectively, in the abomasum than in the small intestine. Gene ontology (GO) terms associated with these 1064 genes were analyzed. Twenty-two GO terms were significantly enriched in genes that were differentially expressed between the two tissues (Table 2). Of these GO terms, biological processes such as regulation of gene

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Table 1 Protein-coding genes uniquely expressed in the bovine abomasum or small intestine. The number denotes normalized read counts per million mapped reads (mean ± sd); locus is expressed as chromosome: gene start-gene end. Ensemble gene ID

Annotation

Locus

Abomasum

ENSBTAG00000007989 ENSBTAG00000040084 ENSBTAG00000003281 ENSBTAG00000038891 ENSBTAG00000009840

Alpha-(1,3)-fucosyltransferase Uncharacterized protein Uncharacterized protein Uncharacterized protein Olfactory receptor, family 2, subfamily AE, member 1

9:54432730-54433809 3:119799067-119799975 3:119810308-119811246 9:88244953-88245726 25:37071831-37072784

0.51 0.11 0.07 0.00 0.00

± ± ± ± ±

0.35 0.07 0.04 0.00 0.00

Small intestine 0.00 0.00 0.00 2.26 0.74

± ± ± ± ±

0.00 0.00 0.00 1.30 0.58

Table 2 Gene ontology (GO) terms enriched in genes with significantly different abundance between the bovine abomasum and small intestine. GO id

GO description

Ratioa

P uncorrected

P bonferroni

P fdr

GO:0022607 GO:0071844 GO:0034622 GO:0034621 GO:0006325 GO:0044427 GO:0005694 GO:0051276 GO:0065003 GO:0043933 GO:0045653 GO:0045638 GO:0000786 GO:0006334 GO:0034728 GO:0032993 GO:0065004 GO:0071824 GO:0060968 GO:0045652 GO:0045637 GO:0006352

Cellular component assembly cellular component assembly at cellular level Cellular macromolecular complex assembly Cellular macromolecular complex subunit organization Chromatin organization Chromosomal part Chromosome Chromosome organization Macromolecular complex assembly Macromolecular complex subunit organization Negative regulation of megakaryocyte differentiation Negative regulation of myeloid cell differentiation Nucleosome Nucleosome assembly Nucleosome organization Protein–DNA complex Protein–DNA complex assembly Protein–DNA complex subunit organization Regulation of gene silencing Regulation of megakaryocyte differentiation Regulation of myeloid cell differentiation Transcription initiation, DNA-dependent

84/724 76/572 66/328 66/362 71/409 72/452 54/197 72/496 74/487 74/530 13/15 18/44 61/136 60/157 60/171 61/150 60/161 60/171 10/18 14/22 19/92 17/64

2.19E−08 2.49E−10 1.08E−10 1.32E−10 2.20E−10 2.48E−10 7.53E−11 1.44E−10 8.99E−11 1.74E−10 5.23E−11 3.37E−11 3.55E−11 9.89E−11 6.55E−11 1.20E−10 4.69E−11 6.55E−11 2.18E−08 5.32E−11 2.82E−06 2.16E−07

9.10E−05 1.03E−06 4.49E−07 5.47E−07 9.14E−07 1.03E−06 3.13E−07 5.99E−07 3.74E−07 7.22E−07 2.17E−07 1.40E−07 1.47E−07 4.11E−07 2.72E−07 4.98E−07 1.95E−07 2.72E−07 9.05E−05 2.21E−07 1.17E−02 8.98E−04

0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0000 0.0080 0.0000

FDR, false discovery rate. a The number of genes assigned to the GO term among the genes with significantly different relative abundance between the abomasum and small intestine/total number of genes in the transcriptome assigned to the same GO term.

silencing (GO:0060968), negative regulation of megakaryocytes differentiation (GO:0045653) and negative regulation of myeloid cell differentiation (GO:0045638) were significantly enriched and may be particularly noteworthy. Further studies are needed to verify the biological implications of these changes and if they are associated with resistance to nematode parasites. 4. Biological pathways affected by nematode infection Comparative transcriptome analysis using high-density microarrays and RNA-seq technology has recently been utilized to understand host–parasite interactions in ruminants (Diez-Tascón et al., 2005; Kadarmideen et al., 2011; Knight et al., 2010; Li and Gasbarre, 2009; Li et al., 2010, 2011a,b). These studies have provided novel insight into gene regulatory networks and biological pathways that are disrupted during nematode infections. This approach has the potential to help us unravel molecular mechanisms underlying protective immunity and host resistance to parasitic infection. While the activation of Th2-linked pathways that enhance intestinal smooth muscle contractility and increase intestinal mucus secretion can protect mice

against N. brasiliensis and, with an activated mucosal mast cell response, Trichinella spiralis, it is insufficient against an initial infection with H. bakeri but protective during a drug-abbreviated secondary challenge infection (Finkelman et al., 2004; Shea-Donohue and Urban, 2004). Similarly, the initial Th2 response in ruminants is generally insufficient to render the host refractory to nematode infections (Gasbarre, 1997; Li et al., 2010). For example, O. ostertagi is able to elicit profound changes in the host immune system, including a rapid and enormous expansion of lymphocytes and lymphoid cells, increased expression of IL-4, and extensive changes in the abomasal tissue and its draining lymph nodes. However, these changes do not transform into a rapid and robust protective immunity (Gasbarre, 1997; Li et al., 2007). Recently, we identified three signaling pathways, the complement system, leukocyte extravasation, and acute phase responses (Li et al., 2010) that are significantly induced by nematode infection in immune cattle. Increased expression of complement components C3, factor B, and factor I in the abomasal mucosa of O. ostertagi-infected cattle at both the mRNA and the protein levels is observed during the development of long-term protective immunity (Li et al., 2010). One of the initiators of local complement activation may be related to secretory IgA and IgM. The infection significantly

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up-regulates the expression of J chain (IGJ), polymeric Ig receptor (PIGR), a receptor responsible for trans-epithelial transport of IgA dimers and IgM pentamers into mucosal secretions, and an IgM-specific receptor (FAIM3), suggesting sustained increases in both synthesis and transport of IgA and IgM during the infection. The elevated levels of inflammatory cytokines, such as IL-4 and IL-1␤, during infection may be involved in gene regulation of complement components. This could be similar to complement activated pathways of localized anaphylaxis seen with certain responses to food allergens (Khodoun et al., 2009). While enhanced tissue repair and mucin secretion in immune animals may be important host immune responses (Rinaldi et al., 2011), local complement activation is responsible for the development of long-term protective immunity and plays a critical role in rendering resistance against O. ostertagi in cattle (Li et al., 2010). The ruminant response to an experimental Cooperia infection differs from that observed with Ostertagia by virtue of temporal shifts in regulatory gene networks and pathways (Li and Gasbarre, 2009). At the early stage of infection at seven dpi, there is increased expression of genes associated with leukocyte homing mechanisms and complement activation. Recent RNA-seq based transcriptome analysis suggested that muscle contraction is the predominant biological process enhanced at this stage of infection (Li and Schroeder, 2012). The primary function of two of the four regulatory networks affected is related to skeletal and muscular systems. Of 34 other pathways significantly affected, the immune pathways related to an acute phase response, leukocyte extravasation, and antigen presentation are activated at seven dpi, consistent with previous findings (Li and Gasbarre, 2009). Other pathways most significantly affected are calcium signaling and actin cytoskeleton signaling. These findings suggest that Cooperia may be able to induce host actin cytoskeleton remodeling and hijack host cell processes to ensure penetration and settlement in the host intestinal epithelium. Furthermore, enhanced smooth muscle contractility at the site of infection may play an important role in host immune responses against a primary C. oncophora infection, as has been shown for resistance in some GI nematode mouse models (Zhao et al., 2008). As the infection progresses, host immune responses shift from muscle contraction and leukocytes migration to increased release of neutrophils, migration of dendritic cells, and natural killer cells. This is similar to localized cellular response in the intestinal sub-mucosa of H. bakeri-infected mice (Anthony et al., 2006). Activation of eosinophil rolling and egression is evident at later days of infections (42 dpi). Other canonical pathways and regulatory networks are significantly altered during the infection as well (Li and Gasbarre, 2009; Li et al., 2011a,b). For example, cholesterol synthesis and retinoic acid synthesis pathways are altered by C. oncophora at seven dpi. As the infection advances to 14 dpi, lipid metabolism is affected, involving SLC27A6 which imports polyunsaturated fatty acid (PUFA) into the host cell. An immune stimulatory role of PUFA in the development of host resistance to infections by C. oncophora and O. ostertagi has been suggested in calves (Muturi et al., 2005). In cattle with previous parasite exposure, re-infection by

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C. oncophora also induced activation of many additional biological pathways (Li et al., 2011b). Notably, the vitamin D receptor activation pathway was strongly induced and may be important in the development of acquired resistance via its potential roles in immune regulation and maintenance of intestinal mucosal integrity. The expression of inducible nitric oxide synthase (NOS2) was also strongly induced during re-infection. As a result, several biological pathways associated with NOS2 are affected. Reactive nitric oxide (NO), produced by NOS and expressed in various cell types, has been shown to control parasitic infection in a murine model (Bogdan, 2001). Indeed, recent studies indicated that host immune responses against helminth infections often involve multiple biological pathways. Higher levels of expression of IgE receptor, integrins, complement, and tissue factors may be related to the manifestation of host resistance in cattle (Araujo et al., 2009). In small ruminants, nematode infection elicited a strong Th2 immune response similar to that in cattle (Hein et al., 2010; Miller and Horohov, 2006; Shakya et al., 2009), including an enhanced production of IL4, IL5, IL13, and IgE antibodies, as well as increased recruitment of eosinophils and mast cells (Meeusen et al., 2005). Goats and sheep are often infected by the same parasite species, provoking similar pathological changes in both host species. However, there is sufficient evolutionary divergence that goats develop a different set of strategies to regulate parasitic infections (Hoste et al., 2008). A direct comparison of FECs from sheep and goats grazing together on the same pasture suggested that the mechanisms of resistance in goats differ from those in sheep, especially with regard to the sequence for establishing immunity (Pomroy et al., 1986). In addition, earlier studies using real-time PCR showed that parasite resistant and susceptible genotypes within the same host species differ in response to related GI parasites like H. contortus and T. colubriformis. Indeed, several studies have confirmed that the host expresses different or unique mechanisms against various parasitic species (Andronicos et al., 2010). Conserved features of a host immune response evoked by resistant sheep include markers of an early inflammatory response, such as Toll-like receptors (TLR2, 4, 9) and free radical producing genes, dual oxidase 1 (DUOX1) and NOS2 (Ingham et al., 2008). Similar responses in humans infected with GI nematode parasites have been associated with altered innate immunity linked to microbial pathogens (Jackson et al., 2006). Recent proteomic and network analyses of genetically resistant and susceptible sheep have identified mitochondrial function and energy partitioning as important components of an effective protective response to nematodes, which operates locally in the abomasal mucosa during the initial interaction between the host and H. contortus (Nagaraj et al., 2012). Microarray-based transcriptome analysis of sheep intestinal immune cells showed down-regulation of immune genes during infection with T. colubriformis. These immune genes affect antigen presentation, caveolarmediated endocytosis, and protein ubiquitination process of the migratory immune cells (Knight et al., 2010). Many transcriptional regulators exert biological function via post-transcriptional mechanisms with subtle or no apparent changes at the level of mRNA expression.

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Therefore, the biological interpretation of high-throughput transcriptome expression data requires both differential expression and differential network analyses (de la Fuente, 2010). The latter relies on powerful computational tools to extract accurate regulatory gene networks reflecting causal interactions underlying biological processes or phenotypes (Baldwin et al., 2012). Recently weighted gene co-expression network analysis (WGCNA) has been used to identify gene modules correlated with the length of infection and numerous hub genes or master regulators in the GI tract of sheep co-infected by H. contortus and T. colubriformis (Kadarmideen et al., 2011). In response to H. contortus infection, regulatory networks that have been identified were associated with T-cell and B-cell regulation, as well as the production of TNF␣, interleukins, and cytokines. On the other hand, regulatory networks associated with T. colubriformis infection are related to protein catabolic process, heat shock protein binding, protein targeting and localization, cytokine receptor binding, TNF receptor binding, apoptosis, and IGF binding. Furthermore, the breeding values of selected hub genes can be applied to breeding strategies to enhance the utility of global gene expression data.

5. Bridging QTL and transcriptome analyses Mounting evidence suggests that utilization of QTLs for applied breeding is particularly beneficial for traits with low heritability (Gibson, 2005), such as parasite resistance. QTL analysis enables investigators to dissect genetic architecture of a trait and to identify underlying candidate genes. A desire to breed for parasite resistant ruminants to reduce dependence on anthelmintics has led to the identification of many putative QTLs that affect FEC and eosinophil counts, especially in small ruminants. Two significant and six suggestive QTLs have been identified in resistant cattle under conditions of natural infection (Coppieters et al., 2009). Significant QTLs associated with FECs are located on two chromosomes, BTA9 and BTA19. A recent report suggested that variation in the susceptibility of H. contortus infection is not within the interferon gamma gene (INFG) but within a QTL segregating with the INFG gene in Rasa Aragonesa ewes (Dervishi et al., 2011). In another study, four putative QTLs are associated with eosinophil counts and FECs in sheep (Dominik et al., 2010). Using a disequilibrium-linkage approach, researchers have identified more refined QTL intervals (<10 cM) in Merion sheep that are related to resistance against infection with H. contortus (Marshall et al., 2009). QTLs on chromosomes 2, 3, and 14 have been identified that are associated with altered FECs from infection with Nematodirus spp. (Crawford et al., 2006). A QTL associated with FECs and eosinophil counts have been located on chromosome 23 adjacent to MHC genes in Australian goats (Bolormaa et al., 2010). The identification of QTLs in small ruminants offer special promise for exploiting marker-assisted selection due to historically high-levels of genetic diversity and strong recent selection in breed formation (Kijas et al., 2012). It is foreseeable that advances in high-throughput genomic technologies will identify causal genes within each QTL.

Progress in genotyping and transcriptome characterization using high-density oligo microarrays and RNA-seq technology has spurred studies of expression quantitative trait loci (eQTL). This approach provides a means to unravel the role of transcriptional regulatory variation beneath complex traits (Michaelson et al., 2009; Montgomery and Dermitzakis, 2011). The power of this approach has been exemplified by a recent study of celiac disease, a heritable chronic inflammatory condition of the small intestine activated by gluten in the diet (Dubois et al., 2010). An eQTL analysis of blood samples revealed a significant correlation between cis gene expression (eQTLs) at 20 of 38 loci associated with significant celiac risk variants. Many of the genetic risk variants were located near genes related to immune function. Similarly, integration of peak eQTL signals and GWA has the potential to identify regulatory pathways and candidate genes involved in resistance to GI nematode parasites in ruminants. Caution is needed in the selected breeding of resistance traits for GI nematode parasites because of the counter-regulatory immune pathways that would predict inappropriate responses to bystander protozoan, bacterial, and viral infections (Brady et al., 1999; Chen et al., 2006; Khan et al., 2008; Noland et al., 2008; Porto et al., 2005). In fact, QTL mapping of resistance to parasites in sheep would argue that selective pressure favors a diverse genome capable of responding to multiple classes of pathogens (Beraldi et al., 2007). A more prudent approach would be the identification of innate resistance traits that can effectively minimize parasite establishment, thereby precluding inflammation and the counter-regulatory pathways inherent in parasitism, and the use of dietary components that directly inhibit parasite development in the host or can transcriptionally induce responses elements associated with host protection (Athanasiadou et al., 2011; deSchoolmeester et al., 2006; Keane et al., 2006). 6. Conclusion Nematode infections in ruminants induce significant changes in patterns of gene expression at the site of infection and in draining lymph nodes particularly for the regulation of cytokines and cell adhesion molecules. Each host–parasite interaction has unique effector mechanisms and associated gene expression profiles. In addition, the host exhibits different immune responses that are temporally regulated during the infection, coinciding with critical changes in parasite development, physiology, and virulence. Understanding the temporal effect of host immune responses during the course of parasitic nematode infections will facilitate the development of optimal strategies to control disease. Moreover, the development of protective immunity and resistance to nematode infections depends on the precise control of expression by the host genome. Regulatory programs controlling the transcriptome evolved at a rapid rate comparable to that of other genomic processes. The identification of regulatory elements that control transcriptome dynamics is crucial to enhance the functional relevance of these pathways as applied to parasite control. Comparative transcriptome analysis in concert with high-throughput genomic

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identification of genome-wide association QTLs affecting host resistance has emerged as a technique to unravel the molecular basis of regulatory networks to explain the complex traits associated with parasite resistance. The challenge is to combine the expanding base of knowledge of parasite products and host resistance mechanisms with a sound strategy of breeding, management, and nutrition to activate inducible pathways that negatively affect GI parasite infection without activating equally harmful unintended consequences. Acknowledgements Mention of trade names or commercial products in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the U. S. Department of Agriculture. The USDA is an equal opportunity provider and employer. References Albers, G.A., Gray, G.D., Piper, L.R., Barker, J.S., Le Jambre, L.F., Barger, I.A., 1987. The genetics of resistance and resilience to Haemonchus contortus infection in young merino sheep. Int. J. Parasitol. 17, 1355–1363. Almería, S., Canals, A., Zarlenga, D.S., Gasbarre, L.C., 1997. Isolation and phenotypic characterization of abomasal mucosal lymphocytes in the course of a primary Ostertagia ostertagi infection in calves. Vet. Immunol. Immunopathol. 57, 87–98. Amarante, A.F., Craig, T., Ramsey, W., El-Sayed, N., Desouki, A., Bazer, F., 1999. Comparison of naturally acquired parasite burdens among Florida Native. Rambouillet and crossbreed ewes. Vet. Parasitol. 85, 61–69. Andronicos, N., Hunt, P., Windon, R., 2010. Expression of genes in gastrointestinal and lymphatic tissues during parasite infection in sheep genetically resistant or susceptible to Trichostrongylus colubriformis and Haemonchus contortus. Int. J. Parasitol. 40, 417–429. Anthony, R.M., Urban Jr., J.F., Alem, F., Hamed, H.A., Rozo, C.T., Boucher, J.L., Van, R.N., Gause, W.C., 2006. Memory T(H)2 cells induce alternatively activated macrophages to mediate protection against nematode parasites. Nat. Med. 12, 955–960. Araujo, R.N., Padilha, T., Zarlenga, D., Sonstegard, T., Connor, E.E., Van Tassel, C., Lima, W.S., Nascimento, E., Gasbarre, L.C., 2009. Use of a candidate gene array to delineate gene expression patterns in cattle selected for resistance or susceptibility to intestinal nematodes. Vet. Parasitol. 162, 106–115. Artis, D., Grencis, R.K., 2008. The intestinal epithelium: sensors to effectors in nematode infection. Mucosal Immunol. 1, 252–264. Artis, D., Wang, M.L., Keilbaugh, S.A., He, W., Brenes, M., Swain, G.P., Knight, P.A., Donaldson, D.D., Lazar, M.A., Miller, H.R., Schad, G.A., Scott, P., Wu, G.D., 2004. RELMbeta/FIZZ2 is a goblet cell-specific immune-effector molecule in the gastrointestinal tract. Proc. Natl. Acad. Sci. U. S. A. 101, 13596–13600. Athanasiadou, S., Jones, L.A., Burgess, S.T., Kyriazakis, I., Pemberton, A.D., Houdijk, J.G., Huntley, J.F., 2011. Genome-wide transcriptomic analysis of intestinal tissue to assess the impact of nutrition and a secondary nematode challenge in lactating rats. PLoS One 6, e20771. Athanasiadou, S., Pemberton, A., Jackson, F., Inglis, N., Miller, H.R., Thévenod, F., Mackellar, A., Huntley, J.F., 2008. Proteomic approach to identify candidate effector molecules during the in vitro immune exclusion of infective Teladorsagia circumcincta in the abomasum of sheep. Vet. Res. 39, 58. Baldwin, R.L., Wu, S., Li, W., Li, C., Bequette, B.J., Li, R.W., 2012. Quantification of transcriptome responses of the rumen epithelium to butyrate infusion using RNA-seq technology. Gene Regul. Syst. Biol. 6, 67–80. Barger, I.A., Gibbs, H.C., 1981. Milk production of cows infected experimentally with trichostrongylid parasites. Vet. Parasitol. 9, 69–73. Bassetto, C.C., Silva, B.F., Newlands, G.F.J., Smith, W.D., Amarante, A.F.T., 2011. Protection of calves against Haemonchus placei and Haemonchus contortus after immunization with gut membrane proteins from H. contortus. Parasite Immunol. 33, 377–381.

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