IAI Accepted Manuscript Posted Online 16 February 2016 Infect. Immun. doi:10.1128/IAI.01309-15 Copyright © 2016 Maaz et al. This is an open-access article distributed under the terms of the Creative Commons Attribution 4.0 International license.
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Susceptibility to ticks and Lyme disease spirochetes is not affected in mice co-infected with nematodes
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Denny Maazac, Sebastian Rauscha, Dania Richterb, Jürgen Krückenc, Anja A. Kühld, Janina Demelerc, Julia Blümkec, Franz-Rainer Matuschkae, Georg v. Samson-Himmelstjernac, Susanne Hartmanna#
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Institute of Immunology, Centre of Infection Medicine, Freie Universität Berlin, Germanya
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Environmental Systems Analysis, Institute of Geoecology, Technical University of Braunschweig,
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Germanyb
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Institute of Parasitology and Tropical Veterinary Medicine, Freie Universität Berlin, Germanyc
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Department of Medicine I for Gastroenterology, Infectious Disease and Rheumatology, Research Center
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ImmunoSciences, Charité Berlin, Germanyd
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Outpatient Clinic, University of Potsdam, Germanye
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Running head: Nematode/tick co-infection
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# Address correspondence to Susanne Hartmann,
[email protected]
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Phone: +49 30-838-51824 Fax: +49 30-838-451824 D.M. and S.R contributed equally to this work.
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Abstract
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Small rodents serve as reservoir hosts for tick-borne pathogens, such as spirochetes causing Lyme disease.
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Whether natural co-infections with other macroparasites alter the success of tick feeding, anti-tick immunity
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and the host’s reservoir competence for tick-borne pathogens remains to be determined. In a parasitological
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survey of wild mice in Berlin approximately 40% of Ixodes ricinus infested animals harboured the nematode
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Heligmosomoides spp. simultaneously. We, therefore, aimed to analyse the immunological impact of the
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nematode/tick co-infection as well as its effect on the tick-borne pathogen Borrelia afzelii. Hosts
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experimentally co-infected with H. polygyrus and larval/nymphal I. ricinus ticks developed substantially
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stronger systemic Th2 responses, based on GATA-3 and IL-13 expression, than did single-infected mice.
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During repeated larval infestations, however, anti-tick Th2 reactivity and an observed partial immunity to
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tick feeding was unaffected by concurrent nematode infections. Importantly, the strong systemic Th2
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immune response in co-infected mice did not affect the susceptibility to tick-borne B. afzelii. An observed
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trend for decreased local and systemic Th1 reactivity against B. afzelii in co-infected mice did not result in
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higher spirochete burden, neither facilitated bacterial dissemination nor induced signs of immunopathology.
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Hence, this study indicates that strong systemic Th2 responses in nematode/tick co-infected house mice do
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not affect the success of tick feeding and the control of the causative agent of Lyme disease.
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Introduction
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Ticks are the most important arthropod vectors of pathogens in temperate regions of the northern
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hemisphere. In particular, ticks of the genus Ixodes transmit several pathogens affecting the health of
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humans and life stock, such as Borrelia spp. causing Lyme disease (LD) or relapsing fever and Rickettsia
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spp. of the spotted fever group causing diseases such as aneruptive fever and tick-borne lymphadenopathy
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[1]. Small rodents serve as reservoir hosts for many tick-borne pathogens and the larval stage of Ixodes
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ricinus, the main vector species in Western and Central Europe, frequently feeds on mice and voles [2]. The
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larval tick stage efficiently acquires the pathogen when feeding on infected rodents [3]. After moulting,
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nymphal ticks constitute the major source for infections in people [4]. Two genospecies of LD spirochetes
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account for the majority of infections in Eurasia, Borrelia garinii and Borrelia afzelii, of which the latter
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mainly and efficiently perpetuates in rodents [5-8]. Mice and voles in urban and rural areas are frequently
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infected by intestinal helminths, such as the trichostrongyloid Heligmosomoides polygyrus [9-11], raising
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the question whether such natural co-infections may affect the feeding success of ticks and the transmission
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of tick-borne pathogens.
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Ticks initiate type 2 T cell (Th2) responses during their blood meal [12]. Although feeding of I. ricinus ticks
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results in protective immunity in guinea pigs and rabbits, animals non-native to this tick’s distribution,
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immune responses are insufficient in protecting most autochthonous wild mouse species and laboratory
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mouse strains against repeated infestations with ticks [12-16]. On the other hand, feeding Ixodes ticks
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suppress Th1 responses against tick-borne LD spirochetes in mice [17], and suppression of Th2 cytokines or
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reconstitution of TNF-α, IL-2 and IFN-γ after tick attachment reduces infection rates in disease-susceptible
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laboratory mouse strains [18, 19]. Infections with the enteric nematode H. polygyrus elicit strong local and
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systemic Th2 and regulatory responses [20, 21] and, thereby, affect the Th1-dependent control of
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concomitant bacterial and protozoan infections [22-28]. It may be that the concurrent presence of enteric
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nematodes synergistically enhances Th2 and associated innate effector cell responses towards repeated tick 3
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infestations and as a result affects tick feeding success and transmission of tick-borne pathogens. The
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expected strong general Th2 skewing of immune responses in mice simultaneously harbouring the two
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macroparasites, H. polygyrus and I. ricinus, may restrain immune control of tick-borne pathogens and
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subsequently increase the reservoir competence of rodents.
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To investigate how frequently mice simultaneously harbour enteric nematodes and ticks in nature, we
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captured wild rodents in Berlin and determined the frequency of infection with Heligmosomoides spp. and
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infestation with I. ricinus. Using a defined experimental laboratory system we then assessed parameters of
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anti-tick immunity and the reservoir competence for LD spirochetes using inbred house mice. To examine
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whether the expected strong Th2 immune responses associated with intestinal nematode infections interfere
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with, first, the experimental host’s development of anti-tick immunity measured my means of tick-specific
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cytokine responses and success of repeated tick feeding, and, second, the control of tick-borne pathogens,
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we established an experimental model of co-infection in inbred laboratory mice, compared susceptibility to
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tick feeding and to tick-borne infection with LD spirochetes and evaluated local and systemic immune
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responses.
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Here we show that (1) Heligmosomoides spp. and I. ricinus frequently parasitize wild Apodemus spp. mice
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in Berlin; (2) local anti-I. ricinus immune responses in laboratory house mice are not altered by a concurrent
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nematode infection, despite the induction of extraordinarily high systemic Th2 responses by nematode/tick
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co-infections; (3) the nematode infection does not alter the feeding success of pathogen-free tick larvae and
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spirochete-infected nymphs as well as the development of partial immunity towards repeated tick
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infestations, and (4) the nematode co-infection does not affect tick-borne transmission, replication and
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dissemination of B. afzelii spirochetes in house mice.
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Material and Methods
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Wild rodent trapping
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Wild rodents were trapped at two urban (Moabit, Steglitz) and two peri-urban study sites (Gatow, Tegel) in
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Berlin, Germany in November 2010 and from April to November 2011 (Landesamt für Gesundheit und
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Soziales, Berlin, Germany, LAGeSo Reg. Nr. G 0210/10). Sites, trapping method and euthanasia were
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described in Krücken & Blümke et al., 2015 (submitted). After animals were transferred to the laboratory,
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the fur was screened for ticks under a stereo microscope. Subsequently, a complete necroscopy was
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performed and the gastro-intestinal tract was thoroughly screened for Heligmosomoides spp., which were
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stored in 80% ethanol. The remaining carcass was placed in screw top jars over water to allow detachment
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of remaining ticks. The water and the rodent body were examined during the following week. Ticks were
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stored in 70% ethanol. Parasite species were determined microscopically according to morphologic literature
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of nematodes [29] and ticks [30-32].
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Laboratory mice, parasites and Borrelia
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Six to eight week old specific pathogen-free (SPF) female C57Bl/6JRj, BALB/cJRj and C3H/HeNRj mice
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(Janvier, France) were used for the experiments. The trichostrongyloid nematode H. polygyrus was
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maintained by serial passage in C57Bl/6 mice. Specific pathogen-free larval I. ricinus ticks were obtained
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from egg batches of two adult females after feeding on laboratory beagle dogs (Landesamt für Gesundheit
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und Soziales, Berlin, Germany, LAGeSo Reg. Nr. H0078/10). Absence of tick-borne pathogens in tick
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larvae used for experimentation was surveyed by published PCRs [33, 34] for a 153bp fragment of the hbb
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gene of Borrelia burgdorferi sensu lato species, a 203bp fragment of the gltA gene of spotted-fever
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Rickettsia spp., a 602-639bp fragment of the 18S rRNA gene of piroplasmida [33], a 257bp fragment of the
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16S rRNA gene of Anaplasmataceae [34] in isolated DNA [33] from subsets of 20 larvae randomly chosen
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from each tick batch. Borrelia afzelii-infected I. ricinus nymphs were derived from larvae that had engorged 5
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on experimentally infected mice (Landesamt für Umwelt, Gesundheit und Verbraucherschutz, Potsdam,
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Germany, licence number 23-2347-A-24-1-2010).
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Animal experimentation
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Mice were infected with 200 H. polygyrus L3 larvae via oral gavage. In an initial experiment, mice of three
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strains (C57Bl/6, BALB/c and C3H) were infested with I. ricinus larvae on day 15 post H. polygyrus
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infection and dissected 20 days later. To survey the feeding success of larval ticks and anti-larval tick
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immune responses, C57Bl/6 mice were infected with H. polygyrus and subsequently repeatedly infested
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with 50 specific pathogen-free I. ricinus larvae on day 4, 18 and 34 p.i. in order to cover the early phase of
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local Th2 reactivity (approx. day 6 p.i.), maximal Th2 reactivity (around day 21 p.i.) and chronic phase of
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infection with declining magnitude of the nematode-induced Th2-reaction (approximately day 35 p.i.) by bi-
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weekly tick infestations. Mice exposed either to nematodes or ticks and naïve mice served as controls. To
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investigate the effect of H. polygyrus on the control of a tick-borne pathogen, nematode-infected and -free
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groups were infested with 4-6 B. afzelii-infected I. ricinus nymphs on day 16 post H. polygyrus infection in
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order to challenge mice with B. afzelii at the peak of nematode-induced Th2-reactivity. Age-matched naïve
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mice served as controls. During the period of tick infestation, all mice including naïve controls wore a collar
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to prevent removal of ticks placed in the head region. Mice were housed individually at 80% humidity in
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filter-topped cages in ventilated cabinets. To optimize recovery of detached ticks, mice were kept on metal
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grids over water and food was restricted to 3g per day during tick feeding. Mice received water ad libitum.
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In several experiments, mice were anesthetized with isoflurane once a week for retroorbital blood collection
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with haematocrit capillary tubes. To assess the feeding success of ticks, I. ricinus larvae or nymphs
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detaching from hosts were collected from the water and counted twice a day, pooled for every mouse in a
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screen-capped glass vial and transferred to a desiccator with supersaturated MgSO4 solution. Feeding
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duration was calculated as mean recovery time of engorged ticks. Seven days after all ticks had detached,
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the engorgement weight of pooled ticks was determined using a micro scale and the mean weight calculated. 6
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Moulting rates of fed larvae from the 1st and 2nd infestation or of fed nymphs were determined 3 months
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after infestation. All animal experiments were approved by and conducted in accordance with guidelines of
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the appropriate committee (Landesamt für Gesundheit und Soziales, Berlin, Germany) under licence number
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LAGeSo Reg. Nr. G0282/12.
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Preparation of nematode, tick and spirochete antigens
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H. polygyrus antigen (Ag) was prepared as described previously [28]. Antigen from feeding larval ticks was
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acquired from specific pathogen-free I. ricinus larvae 48 h after attachment to C57Bl/6 mice, during
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intensive saliva production. Ticks were removed from the host, washed in sterile PBS, homogenised and
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sonicated (3×10 s, 60W) in PBS on ice and centrifuged at 20,000×g for 10 min at 4°C. The supernatant was
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sterile-filtered through 0.22µm filters (Millipore, USA). For B. afzelii antigen, spirochetes cultured in BSK-
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H medium (Sigma-Aldrich, Germany) supplemented with 6% rabbit serum were washed three times by
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centrifugation at 10,000×g at 4°C for 30 min in PBS containing 5mM MgCl2, sonicated on ice (8×15 s) and
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centrifuged at 10,000×g at 4°C for 30 min. Protein contents of the supernatants were determined by BCA
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protein assay kit (Pierce, USA). Antigens were stored at -80°C until use.
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Cell culture
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Splenocytes and cells from gut-draining and head-skin-draining lymph nodes were isolated from euthanized
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mice by passing the organs through a 70µm cell strainer (BD, USA). Cells were cultured as triplicates of
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3.5-5×105 cells in 200µl RPMI 1640 (PanBiotech, Germany) containing 10% fetal calf serum, 20mM L-
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glutamine, 100U/ml penicillin and 100µg/ml streptomycin. Cultures were stimulated with 40µg/ml worm or
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tick Ag, 30µg/ml B. afzelii Ag (all for 72 h) or with 1µg/ml αCD3 and αCD28 (BD, USA) for 48 h at 37°C
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and 5% CO2. Supernatants were stored at -20°C for cytokine detection.
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Immunohistology
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Formalin fixed tissue samples were dehydrated and embedded in paraffin. Tibiotarsal joints were decalcified
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prior to paraffin embedding. 1-2µm paraffin sections were de-waxed and hydrated and histochemically
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stained for overview with hematoxylin and eosin (Merck), for mast cells using toluidine blue and for
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eosinophil granulocytes employing a modified sirius red staining protocol [35]. Sections were stained with
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toluidine blue (Sigma) for 10 min, washed with distilled water and dehydrated by dipping in 1% acetic acid
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and 100% ethanol. Sections were cleared in xylol and coversliped with corbit balsam (Hecht). For staining
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of eosinophils, sections were stained for 1 min with hematoxylin (Merck), washed with tap water and
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dehydrated with 100% ethanol before staining with direct red 80 (Sigma) for 2 h. Sections were washed with
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tap water, dehydrated with 100% ethanol and cleared in xylol before coverslipping with corbit balsam.
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Images were acquired using a fluorescence microscope (AxioImager Z1) equipped with a CCD camera
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(AxioCam MRm) and processed with Axiovision software (Carl Zeiss AG, Germany).
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ELISA
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Culture supernatants were screened for IFN-γ, IL-13 (Ready-Set-Go! ELISA, eBioscience, USA) and IL-10
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(BD, USA) according to the manufacturer’s instructions.
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Flow cytometry
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Blood was sampled in PBS with 0.2% BSA and 5mM EDTA and cells were stained with αCD4-FITC
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(RM4-5, eBioscience, USA) and fixable viability dye-eFluor780 (eBioscience, USA) for dead cell
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exclusion. Red blood cells were lysed in FACS Lysing Solution (BD, USA) before leucocytes were stained
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intracellularly with αFoxp3-eFluor450 (FJK-16s, eBioscience, USA), αGATA-3-eFluor660 (TWAJ,
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eBioscience, USA) and αT-bet-PE (eBio4B10, eBioscience, USA) using a fixation/permeabilization buffer
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kit (eBioscience, USA). Cells from spleen and lymph nodes were labelled with αCD4-PerCP-eFluor710 8
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(GK1.5, eBioscience, USA) and fixable viability dye-eFluor780 (eBioscience, USA) in PBS with 0.2%
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BSA. Transcription factors were stained according to white blood cells. For intracellular detection of
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cytokines, cells were incubated with 1µg/ml PMA and ionomycin for 30 min and 5µg/ml brefeldin A
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(Sigma, Germany) for another 2.5 h. Subsequently, they were stained with αIFNγ-eFluor450 (XMG1.2,
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eBioscience, USA), αIL-13-AlexaFluor647 (eBio13A, eBioscience, USA), αIL-10-PE (JES5-16E3,
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eBioscience, USA) and αIL-17-AlexaFluor488 (TC11-18H10, BD, USA). Cells were acquired using LSRII
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and Canto II flow cytometers (BD, USA) and analysed using FloJo 8.8.7 software (TreeStar, USA).
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Spirochete quantification
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For quantification of spirochetes in mouse organs in relation to tissue weight, a touchdown-qPCR assay was
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developed for absolute quantification with λ phage genome copies as an external standard. For DNA
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isolation, 80 ±1mg of head skin and 70 ±1mg of heart were sampled with sterile scalpel blades, the skinned
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left tibiotarsal joint (range 32.6-39.1mg) was obtained using clean scissors. Prior to DNA isolation with the
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FastDNA Spin Kit with Lysing Matrix A (MPBiomedicals, France), 104 copies of 48502bp λ genome
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(Thermo Fisher Scientific, USA) per mg head skin or heart and 105 copies per tibiotarsal joint were added as
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an external standard for tissue weight. To calculate copy numbers, the DNA concentration (0.3µg/µl
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measured spectrophotometrically by the company) and the molecular weight of the genome were used.
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Samples were homogenised with the FastPrep 24 (MP Biomedicals, France) for 4×30 s with 6.0m/s and
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cooled on ice between homogenisation steps. Variations of the DNA amount in the samples used in qPCR
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have a major impact on the accuracy of the absolute quantification, since qPCR efficiency is decreased with
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higher DNA concentrations. As common spectrophotometrical DNA quantification is strongly impacted by
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contaminations with salts and proteins, a more precise fluoroscopical DNA quantification assay was
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developed using the LCGreen Plus 10× dye (BioChem, USA), exclusively binding on dsDNA and the
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LightCycler480 II System (Roche Molecular Diagnostics, USA) for fluorescence measurement. Accurate
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DNA quantification in a range of 1 to 100ng was obtained. On 96-well plates, DNA samples, standards (20, 9
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40, 60, 80 and 100ng λ phage DNA) and water controls were prepared in duplicates in 10µl assay volume.
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After vigorous vortexing of sealed plates, samples were centrifuged and fluorescence was measured by the
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LightCycler480 II after 2 min incubation at 25°C during short heating from 25 to 26°C with 100 acquisitions
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per °C. For extrapolation of dsDNA-quantity in samples, a calibration curve was calculated from
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fluorescence values of the standards using a 4-parameter logistic regression model. Two real-time PCR
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assays targeting a 153bp fragment of the hbb gene of LD spirochetes and a 158 bp fragment of λ phage
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genome were conducted with the LightCycler480 II using LightCycler 480 SYBR Green I Master (Roche
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Molecular Diagnostics, USA), 0.5µM of each primer and exactly 100ng sample DNA in a 20µl volume.
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Primers used for hbb fragment are published elsewhere [36], forward and reverse primer used for λ phage
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genome were 5'-TTTTGATTTTGCTGTTTCAA-3' and 5'-ACCTTTTCCATGAATTGGTA-3'. All PCR
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reactions were performed as duplicates and included a negative control and standards, each with 100ng
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target-free mouse DNA of the respective organ. Standards consisted of a dilution series of 104-101 and 3
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copies of an hbb plasmid standard or λ phage DNA. The thermocycling conditions consisted of an initial
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denaturation step at 95°C for 5 min followed by 50 cycles of 95°C for 5 s, annealing for 30 s (initial
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temperature 62°C, then decreasing by 0.25°C/cycle until 57°C) and 72°C for 15 s. Touchdown amplification
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was performed to increase binding specificity of primers during the first cycles. After amplification, a high
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resolution melting step consisting of 95°C for 5 s, 40°C for 1 min and progressive heating from 65 to 95°C
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with 20 acquisitions per °C was performed to identify target fragments by melting temperature in
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combination with their fragment size in subsequent agarose gel electrophoresis. Borrelia hbb copies
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extrapolated from the standard curve were multiplied with the ratio of extrapolated λ phage DNA copies in
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the sample and number of copies previously added per mg skin/heart or per tibiotarsal joint, respectively.
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The detection limit was less than 3 copies and the quantification limit 5-10 copies per 100ng DNA for both
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touchdown-qPCR assays.
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Statistics 10
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Confidence intervals (CI) of prevalences were calculated as Wilson score intervals using R Statistics and the
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“PlotCIs” package. Negative binomial regression of tick numbers was performed using R Statistics Software
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and “MASS” package to estimate the influence of the variable Heligmosomoides spp. count and categorical
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variables mouse species (A. flavicollis and A. sylvaticus) and trapping location (Gatow, Moabit, Steglitz and
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Tegel) on the number of I. ricinus ticks on wild mice.
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GraphPad Prism 5.01 (San Diego, USA) was used for plots and statistical analysis of experimental infection
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data. Statistical differences were tested using unpaired t-test, one-way ANOVA with Bonferroni-corrected
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post-test (Fig. 3 and 5) and two-way-ANOVA (Fig. 2) under the assumption of normal distribution of FACS
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fluorescence data, ELISA optic density data and the majority of the experimental parasite infection data. The
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post-tests following two-way-ANOVA for individual pairs using the Bonferroni correction were performed
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with the “Post test calculator” at http://graphpad.com/quickcalcs. The number of engorged tick nymphs and
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moulting rate in Fig. 7A, D and cell counts in Fig. S1 B were not normally distributed and differences were
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tested using the Mann-Whitney-U-Test. Levels of significance below p=0.05 were considered to be
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significant.
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Results
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Nematode/tick co-infection is frequent in urban and peri-urban wild mice
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A total of 107 potential murine hosts of Heligmosomoides spp., representing 82 Yellow-necked mice
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(Apodemus flavicollis) and 25 long-tailed wood mice (A. sylvaticus), were live-trapped at 2 urban (Moabit,
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Steglitz) and 2 peri-urban (Gatow, Tegel) sites in Berlin and examined for their parasite burden. No house
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mice (Mus musculus) were caught as traps were placed exclusively outside buildings. Both species
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frequently hosted Heligmosomoides spp. (47.7%, 95% CI 38.5-57.0, N=107) with intensities of 1-246
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specimens (Table 1). In addition, 55.1% (95% CI 45.7-64.2) were infested by up to 59 subadult I. ricinus
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ticks (92.1% larvae, 7.9% nymphs). A high percentage (41.3%, 95% CI 29.1-53.4) of tick-infested mice
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(N=59) were simultaneously infected by Heligmosomoides spp. A negative binomial regression model
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including all 107 mice showed an important influence of the trapping location and/or the rodent species on
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the number of I. ricinus ticks on the rodents but could not distinguish between both variables because of
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total collinearity (Fig. 1). A. sylvaticus mice were exclusively trapped in Moabit whereas A. flavicollis only
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occurred at the three other locations. A. flavicollis from Gatow were infested with significantly more ticks
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than those from Steglitz (p
