The host range of the male-killing symbiont Arsenophonus nasoniae in filth fly parasitioids

Journal of Invertebrate Pathology 106 (2011) 371–379

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The host range of the male-killing symbiont Arsenophonus nasoniae in filth fly parasitioids Graeme P. Taylor a,⇑, Paul C. Coghlin b, Kevin D. Floate b, Steve J. Perlman a a b

Dept. of Biology, U. Victoria, P.O. Box 3020, STN CSC, Victoria, British Columbia, Canada V8W3N5 Agriculture and Agri-Food Canada, P.O. Box 3000, Lethbridge Research Centre, Lethbridge, AB, Canada T1J 4B1

a r t i c l e

i n f o

Article history: Received 10 July 2010 Accepted 4 December 2010 Available online 13 December 2010 Keywords: Arsenophonus Nasonia Male-killer Reproductive parasite Wolbachia Phage

a b s t r a c t The Son-killer bacterium, Arsenophonus nasoniae, infects Nasonia vitripennis (Hymenoptera: Pteromalidae), a parasitic wasp that attacks filth flies. This gammaproteobacterium kills a substantial amount of male embryos produced by an infected female. Aside from male death, the bacterium does not measurably affect the host, and how it is maintained in the host population is unknown. Interestingly, this bacterial symbiont can be transmitted both vertically (from mother to offspring) and horizontally (to unrelated Nasonia wasps developing in the same fly host). This latter mode may allow the bacterium to spread throughout the ecological community of filth flies and their parasitoids, and to colonize novel species, as well as permit its long-term persistence. We tested 11 species of filth flies and 25 species of their associated parasitoids (representing 28 populations from 16 countries) using diagnostic PCR to assess the bacterium’s actual host range. In addition to 16S rRNA, two loci were targeted: the housekeeping gene infB, and a sequence with high homology to a DNA polymerase gene from a lysogenic phage previously identified from other insect symbionts. We identified infections of A. nasoniae in four species of parasitoids, representing three taxonomic families. Highly similar phage sequences were also identified in three of the four species. These results identify the symbiont as a generalist, rather than a specialist restricted solely to species of Nasonia, and also that horizontal transmission may play an important role in its maintenance. Crown Copyright Ó 2010 Published by Elsevier Inc. All rights reserved.

1. Introduction Insects are commonly infected with microbial symbionts that hinder their host’s fitness (i.e., are parasitic), increase their host’s fitness (i.e., are beneficial), or exist between these two extremes. Many symbionts are transmitted in a primarily vertical fashion by their insect host, often via the cytoplasm of the egg. This maternal transmission forms a link between the fitness of the host and the symbiont (Werren and O’Neill, 1997). Selection will typically maintain beneficial symbionts within the host population while eliminating more costly symbionts (Hurst, 1991). To this end, symbionts often provide nutrients or protection to their host (Baumann et al., 1995; Montllor et al., 2002; Oliver et al., 2005). Alternatively, some symbionts manipulate their host’s reproduction to be maintained in the host population (Stouthamer et al., 1999). Maternally inherited symbionts are only transmitted by females, causing male hosts to act as an evolutionary dead-end for the symbiont. The symbiont will thus increase its fitness if it causes an infected host to produce more, or higher quality, daughters than an uninfected host. These alterations to the host’s reproduction oc⇑ Corresponding author. E-mail address: [email protected] (G.P. Taylor).

cur by the symbiont either inducing cytoplasmic incompatibility, or by altering the sex-ratio of the offspring produced. Sex-ratio alterations can occur in either the primary sex-ratio, i.e., feminization or induction of parthenogenesis, or in the secondary sex-ratio of the host, i.e., killing males (Werren et al., 2008). Symbionts that kill males occur in a diverse array of arthropods and can affect several different sex-determination mechanisms (Andreadis, 1985; Hurst, 1991; Hurst and Jiggins, 2000; Nakanishi et al., 2008). Almost all of these agents kill males during the embryonic stage, when it is predicted that (infected) female siblings will benefit the most from male death (Hurst, 1991). These proposed benefits include reduced inbreeding, reduced competition by siblings for local resources, and increased resources through consumption of dead male siblings (Hurst and Jiggins, 2000). Although these benefits are thought to play a crucial role in the maintenance of most male-killers (Hurst et al., 1997), only in one system has a fitness benefit been empirically identified; female pseudoscorpions infected by a male-killing strain of Wolbachia produce daughters of both higher quality and number than uninfected females (Koop et al., 2009). How male-killing symbionts are maintained in other systems remains a mystery. Male-killers that do not benefit their host may rely on horizontal transmission to be maintained. A microsporidium in mosquitoes

0022-2011/$ - see front matter Crown Copyright Ó 2010 Published by Elsevier Inc. All rights reserved. doi:10.1016/j.jip.2010.12.004

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kills males as larvae rather than as embryos (Andreadis, 1985). Upon male death the parasite is released to the environment where it infects additional hosts, and this horizontal transmission is thought to play a crucial role in the maintenance of the parasite (Andreadis, 1985). Although embryonic male-killers are thought to be transmitted primarily maternally, horizontal transmission may augment their successful transmission. Horizontal transmission may play a critical role in maintaining the male-killing bacterium Arsenophonus nasoniae (Gammaproteobacteria: Enterobacteriaceae) in its host Nasonia vitripennis (Hymenoptera: Pteromalidae). This wasp is a common pupal parasitoid of flies found in bird nests (e.g. Protocalliphora sialia), and an occasional parasitoid of other vertebrate-associated filth flies, for example, those that breed in dung or carrion (Floate et al., 1999; Peters and Abraham, 2010). Also known as Son-killer, A. nasoniae inhibits the formation of the maternal centrosome in N. vitripennis embryos and causes 80% of sons to die (Balas et al., 1996; Ferree et al., 2008; Skinner, 1985). The bacterium is found systemically in the bodies of infected females and surviving males without causing apparent effects (either beneficial or detrimental) to the host (Balas et al., 1996; Huger et al., 1985). Wasps infected with A. nasoniae have been found across the continental United States with a prevalence of 4–10% (Balas et al., 1996; Skinner, 1985). Within infected lineages, Son-killer is transmitted with high efficiency (95%) from the female to her offspring in an atypical fashion (Skinner, 1985). Symbiont transmission occurs not through the egg, but via the tissue of the wasp’s fly host which is mechanically inoculated with bacteria by the female wasp during parasitism (Huger et al., 1985). Wasp larvae acquire Son-killer per-orally and are infected through the midgut (Huger et al., 1985; Skinner, 1985). Interestingly, and perhaps related to this mode of transmission, A. nasoniae is unusual for insect symbionts in that it can be easily cultured outside of the host (Gherna et al., 1991; Werren et al., 1986). This ability of the bacterium to live outside cells may allow it to exploit a wide range of hosts. In the field, multiple female Nasonia wasps can parasitize the same fly pupa; i.e., ‘superparasitism’ (Grillenberger et al., 2008). Under these conditions, all wasp larvae within the fly will interact with the bacteria deposited by an infected female. About 95% of the Nasonia emerging from an inoculated host are infected, regardless of their mother’s infection status (Skinner, 1985). Many parasitoid wasp species share overlapping host ranges and can co-parasitize a single host (Floate et al., 1999; Floate, 2002). A sympatric congener, Nasonia longicornis, is also known to be infected with A. nasoniae in the field (Balas et al., 1996). A recent survey also found A. nasoniae in Pachycrepoideus vindemmiae, Muscidifurax raptor, and Spalangia cameroni (Hymenoptera: Pteromalidae), and Protocalliphora azurea (Diptera: Calliphoridae) (Duron et al., 2010). This study also showed in experimental wasp co-infections that A. nasoniae can be horizontally transferred from N. vitripennis to N. longicornis, Nasonia giraulti, and Muscidifurax raptorellus, and that it causes male-killing in these species (Duron et al., 2010). In this study we assessed the host distribution of Arsenophonus in a community of filth flies and their parasitoid wasps from a wide geographic range. This was done using diagnostic polymerase chain reaction (PCR) and targeting the multi-copy 16S and 23S rDNA loci. A single-copy bacterial gene, infB, has previously been used to resolve the Enterobacteriaceae at a species level (Hedegaard et al., 1999). The infB gene encodes a protein essential for initiating protein synthesis, and here we use this locus to determine the relatedness of Arsenophonus strains found in this community. We also screened Arsenophonus-positive insects for a phage gene that has been found in a number of related insect symbionts.

2. Methods 2.1. Sample collection We screened adults from a total of 25 species of Hymenoptera (438 total individuals) and 11 species of Diptera (17 total individuals) for Arsenophonus infection. The majority of these samples were used in a previous screening survey for Wolbachia (Floate et al., 2006, 2008; Kyei-Poku et al., 2006), and represent individuals collected from both laboratory colonies and the field (Table 1). We also obtained additional insects by collecting mountain bluebird (Sialia currucoides) nest material from nest boxes near Lethbridge, AB, on two separate occasions in July, 2008. The nest material was placed within a culture cage incubated at 26 °C (16:8 h light:dark cycle) and emergent insects were identified using a light microscope. All insects were individually washed prior to DNA extraction to remove surface contaminants (1 min with 95% EtOH followed by three rinses of sterile dH2O for 1 min each). DNA was extracted using either the Qiagen Blood and Tissue kit, or by the STE method: the insect was ground using a pipette tip in 25 lL of STE buffer (10 mM Tris buffer at pH 8.0, 1 mM EDTA at pH 8.0, 100 mM NaCl) and 5 lL of proteinase K (20 mg/mL), and incubated at 37 °C for 1 h, followed by 3 min at 96 °C. All DNA was stored at 20 °C until use. We also isolated and cultured A. nasoniae from the N. vitripennis collected from the bluebird nest material. Under sterile conditions, 26 live wasps were individually washed and decapitated using a razor blade. DNA was extracted from the head of the wasp and used to determine infection status. The wasp body was ground in 50 lL of tryptic soy broth (BACTO) and 30 lL was spread onto GC agar supplemented with Kellogg’s supplement B (DIFCO; Kellogg et al., 1963). Plates were incubated at 26 °C for 96 h. Using morphological characteristics (Gherna et al., 1991), A. nasoniae colonies were subcultured from infected wasps. Wasps used for bacterial isolation were not included in the survey results. We also used PCR and sequencing to confirm that the cultured isolates were A. nasoniae (see below).

2.2. Diagnostic PCR screening Infection status for Arsenophonus was initially assessed by amplifying 23S rDNA from insect samples. For samples displaying a positive band, the 16S locus was subsequently amplified and sequenced. Another locus, infB, was amplified and sequenced from the A. nasoniae type strain using degenerate primers. Primers were developed for this locus using Primer3 (Rozen and Skaletsky, 2000) to maximize the mismatches between A. nasoniae and closely related bacteria. These novel primers were used to amplify and sequence the infB locus from Arsenophonus-positive samples. Finally, during the course of this project we identified a sequence in the type strain of A. nasoniae highly similar to the DNA polymerase P45 gene from lysogenic bacteriophage APSE, originally identified in the pea aphid symbiont Hamiltonella defensa. It is not known if the sequence we identified is associated with active phage or is stably inherited in A. nasoniae. It is also possible that this sequence is associated with symbionts other than A. nasoniae (see below). We examined the distribution and evolution of this phage-associated sequence as a window into further understanding the population biology of A. nasoniae. As a positive control for our DNA extractions, we amplified a fragment of the low copy insect gene, histone H3, from DNA samples testing negative in all other reactions. PCR conditions and all primer sequences are listed in Table 2. In all sets of reactions, an appropriate positive control (DNA extracted from A. nasoniae from the American Type Culture Collection [ATCC strain 49151], or a previous DNA extraction known to be

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Table 1 Arsenophonus screening results for a sample of filth flies and their parasitic wasps. Numbers represent infected individuals out of the total number tested for A. nasoniae and APSE P45. Only A. nasoniae infected samples were examined for APSE P45. Insecta

Family

Sourceb

Sexc

Flies (Diptera) Haematobia irritans Lucilla cuprina Musca domestica Neobellieria Phormia sp. Pollenia rudius Protocalliphora sialia

A. nasoniae

P45

Muscidae Calliphoridae Muscidae Sarchophagidae Calliphoridae Calliphoridae Calliphoridae

Lethbridge, AB Bali, Indonesia Lethbridge, AB (lc) AAFC (lc) San Antonio, TX (lc) Lethbridge, AB Lethbridge, AB

2 1 1 1 1 3 3

$ unkn $ unkn unkn unkn $

– – – – – – –

n.td n.t n.t n.t n.t n.t n.t

Protophormia terraenovae Ravinia querula Sarcophaga bullata Sepsis sp.

Calliphoridae Sarcophagidae Sarcophagidae Sepsidae

Lethbridge, Lethbridge, Lethbridge, Lethbridge,

1 2 1 1

unkn unkn unkn unkn

– – – –

n.t n.t n.t n.t

Wasps (Hymenoptera) Aphaereta pallipes Brachymeria podagrica Diapria conica Dirhinus himalayanus

Braconidae Chalcididae Diapriidae Chalcididae

Ottawa, ON Buenos Aires, ARG Lethbridge, AB Morocco

1 1 1 1

$ $ $ $

– – – –

n.t n.t n.t n.t

Eupelmus vesicularis

Eupelmidae

Agassiz, BC Lethbridge, AB Romania Turkey

8 8 4 1

$ $ $ $

1/8 – 2/4 1/1

1/1 n.t 1/2 0/1

Kleidotoma sp.

Eucoilidae

Lethbridge, AB

1

$

–

n.t

Muscidifurax raptor

Pteromalidae

Denmark Lethbridge, AB (lc) Ottawa, ON PNE (lc)

1 8 1 2

$ $ $ $

– – – –

n.t n.t n.t n.t

Muscidifurax raptorellus

Pteromalidae

AAFC (lc) PNE (lc)

1 1

$ $

– –

n.t n.t

Muscidifurax uniraptor Muscidifurax zaraptor

Pteromalidae Pteromalidae

MLIRL (lc) KSU (lc) Lethbridge, AB (lc) PNE (lc)

1 1 2 1

$ $ $ $

– – – –

n.t n.t n.t n.t

Nasonia vitripennis

Pteromalidae

Buenos Aires, ARG Indiana (lc) Lethbridge, AB Lethbridge, AB (lc) Ottawa, ON Utah (lc)

1 31 81 39 1 17

$ $ $ $ $ $

– – 38/81e – – –

n.t n.t 10/11f n.t n.t n.t

Pachycrepoideus vindemiae

Pteromalidae

Denmark AAFC (lc)

1 4

$ $

– –

n.t n.t

Phygadeuon sp. Phygadeuon fumator

Ichneumonidae Ichneumonidae

Denmark Denmark Ottawa, ON

1 3 1

$ $ $

– – –

n.t n.t n.t

Pteromalus venustus Spalangia cameroni

Pteromalidae Pteromalidae

Lethbridge, AB AAFC (lc) Arkansas Denmark Florida France Kazakhstan Mead, Nebraska Minnesota Ottawa, ON Peru Russia

5 39 1 6 7 1 1 1 1 1 2 1

4$1# $ $ $ $ $ $ $ $ $ $ $

– – – – – – – – – – 2/2 1/1

n.t n.t n.t n.t n.t n.t n.t n.t n.t n.t 2/2 1/1

Spalangia endius

Pteromalidae

Arkansas Florida Kazakhstan Maine Mead, Nebraska Minnesota N Illinois University Ottawa, ON Peru PNE (lc) Russia

2 10 8 1 7 7 1 1 1 2 6

$ $ $ $ $ $ $ $ $ $ $

– – – – – 7/7e – – – – –

n.t n.t n.t n.t n.t 3/4f n.t n.t n.t n.t n.t

Spalangia gemina

Pteromalidae

Brazil

4

$

–

n.t

Number screened

AB AB AB AB

(continued on next page)

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Table 1 (continued) Insecta

Family

Sourceb

Number screened

Sexc

A. nasoniae

P45

PNE (lc)

2

$

–

n.t

Spalangia nigra

Pteromalidae

Mead, Nebraska Ottawa, ON UMN (lc)

1 2 1

$ $ $

– – –

n.t n.t n.t

Spalangia nigroaenea

Pteromalidae

Florida Kazakhstan Mead, Nebraska Ottawa, ON Russia UMN (lc)

1 2 11 1 2 1

$ $ $ $ $ $

– – – – – –

n.t n.t n.t n.t n.t n.t

Tachinophaegus zealandicus

Encyrtidae

Buenos Aires, ARG Brazil

1 1

$ $

– –

n.t n.t

Trichomalopsis sarcophagae

Pteromalidae

Lethbridge, AB Lethbridge, AB (lc)

20 20

$ $

– –

n.t n.t

Trichomalopsis viridiscens Trichopria nigra

Pteromalidae Diapriidae

Ottawa, ON Kazakhstan Russia

1 2 2

$ $ $

– – –

n.t n.t n.t

Urolepis rufipes

Pteromalidae

Denmark Lethbridge, AB (lc) Ottawa, ON PNE (lc)

2 20 2 2

$ $ $ $

– – – –

n.t n.t n.t n.t

a Most insect samples were previously used in a screening survey for Wolbachia; see (Floate et al., 2006, 2008; Kyei-Poku et al., 2006) for further information regarding insect host range and food sources. Eupelmidae were not included in the aforementioned study; Eupelmus vesicularis is a generalist parasitoid and hyper-parasitoid (Gibson, 1990), and has been rarely collected from filth fly pupae (Floate et al., 2000; Peters and Abraham, 2010). b lc denotes a laboratory colony, all others refer to location of field collections. If the original collection location of a laboratory culture is unknown, the supplier is listed instead: PNE = PNE, Inc., North Ridgeville, Ohio; UMN = University of Minnesota, KSU = Kentucky State University, AAFC = Agriculture and Agri-Food Canada, MLIRL = Midwest Livestock Insects Research Laboratory. c If no number is given, sex listed is for all tested samples. If present, the number gives individuals of each sex tested. unkn signifies insects were not sexed prior to DNA extraction. d n.t represents samples not tested. e At least two individuals were confirmed infected by direct sequencing of 16S rDNA. The remaining specimens were identified as infected by presence of the expected band. f Only a subset of Arsenophonus samples were screened for P45 sequence; 11 and 4 infected samples of N. vitripennis and S. endius, respectively. Products from 2 N. vitripennis and none from S. endius were confirmed by direct sequencing.

infected) and negative water blank were used. PCR products were visualized on an agarose gel using ethidium bromide staining, and purified for sequencing using a Qiagen Gel Extraction kit or a Qiagen PCR Purification kit. Products were sequenced by Macrogen, South Korea, in both directions using the same primers as were used for the initial amplification unless otherwise noted. Sequences and chromatograms were analyzed, and vector sequences removed if necessary, using Biomatters’ Geneious software. Sequences were identified through BLAST using the top hit by identity. All sequences have been deposited in GenBank under the accessions HM594692-HM594710. 2.3. Screening for alternative phage hosts Because APSE phage sequences are known from other insect symbionts (Hansen et al., 2007; van der Wilk et al., 1999), we wanted to screen for potential bacterial hosts other than A. nasoniae in our insect samples. We performed a preliminary survey using SuPER (suicide polymerase endonuclease restriction) PCR for symbionts other than Arsenophonus within an APSE phageinfected sample. This method removes target DNA (e.g., Arsenophonus) from a sample using restriction digestion (Green and Minz, 2005). Universal primers are subsequently used to amplify the DNA from the remaining bacterial species, which is then cloned and sequenced for identification. One S. cameroni wasp that tested positive for both Arsenophonus and APSE phage was analyzed using SuPER PCR. Kill primers (Table 2), specific for the 16S rDNA of A. nasoniae, were designed using Primer3 (Rozen and Skaletsky, 2000) and used in a reaction mixture containing 1.5 U of Taq DNA polymerase, 10 lg of BSA, 0.86x PCR buffer, 5.6 mM MgCl2, and 0.56 lM of each primer in a 29 lL final reaction volume was incubated at 95 °C for 3 min. The heat was then reduced to 70 °C,

and deoxynucleoside triphosphate (0.172 mM final concentration in the reaction mixture) and 5 U of the restriction enzyme Tsp509I (New England Biolabs, Beverly, MA) along with 1.38x NEBuffer 1, preheated to 70 °C, was added to the sample. This thermostable restriction enzyme removes the annealed sequences from subsequent reactions. The reaction mixture was incubated at 60 °C for 60 min, 95 °C for 30 min, then held at 4 °C, at which time proteinase K (20 mg/mL) was added to a final concentration of 3.3 lg/lL. The mixture was incubated at 58 °C for 30 min, 95 °C for 10 min, then cooled and stored at 4 °C for a subsequent PCR reaction. The reaction mixture was used directly as template in a PCR using universal 16S rDNA primers in a mixture containing 10 lg BSA, 0.96x PCR buffer, 1.92 mM MgCl2, 0.192 mM dNTP, and 1.5 U of Taq. The amplified product was cleaned using a Qiagen PCR Purification kit and a clone library was developed using a Stratogene Cloning Kit. A Qiagen Miniprep Kit was used to prepare 13 clones and these were sequenced using M13 primers (Table 2). All 13 clones sequenced from the SuPER PCR assay shared 99% nucleotide similarity to one another and >99% similarity in BLAST searches to bacteria allied with the insect symbiont Sodalis. To determine whether the Sodalis-like symbiont was present in other APSE phage-positive insects, we designed specific 16S rDNA primers (Table 2) using Primer3 (Rozen and Skaletsky, 2000) and conducted diagnostic PCR. Alongside the APSE phage-positive samples we used a Sodalis-positive control, and three negative controls: DNA extracted from Escherichia coli, DNA extracted from the A. nasoniae type strain, and a water blank. 2.4. Phylogenetic analysis Phylogenetic analyses were conducted on both the infB and viral P45 genes. The infB dataset contains GenBank sequences from

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375

Table 2 PCR primers and cycling conditions. Gene 16S (Arsenophonus)

Primers

Reaction conditions

0

ArsF 5 -GGGTTGTAAAGTACTTTCAGTCGT-3

0

0

ArsR3 5 -CCTYTATCTCTAAAGGMTTCGCTGGATG-3 (Duron et al., 2008)

2 min @ 95 °C; 35 cycles of 30s @ 94 °C, 30s @ 54 °C, 60s min @ 72 °C; 5 min @ 72 °C 0

16S (kill primers)

Ars315F 50 -TCAGTCGTGAGGAAGGTGTTAAGG-30 Ars718R 50 -TGACCACAACCTCCAAATCGACA-30 (This study)

Non-standard cycling conditions, see methods

16S (universal)

63F 50 -CAGGCCTAACACATGCAAGTC-30

4 min @ 94 °C; 38 cycles of 60s @ 95 °C, 60s @ 50 °C, 60s @ 72 °C; 5 min @ 72 °C

907R 50 -CCGTCAATTCCTTTRAGTTT-30 (Marchesi et al., 1998; Schabereiter-Gurtner et al., 2003) 16S (Sodalis)

Sod320F 50 -ATATTGCACAATGGGGGAAA-30

5 min @ 95 °C; 30 cycles of 60s @ 94 °C, 60s @ 56 °C, 60s @ 72 °C; 10 min @ 72 °C

Sod801R 50 -CAAGGCCACAACCTTCAAAT-30 (This study) 23S (Arsenophonus)

Ars23SF 50 -CGTTTGATGAATTCATAGTCAAA-30

5 min @ 95 °C; 34 cycles of 30s @ 94 °C, 30s @ 54 °C, 45s @ 72 °C; 10 min @ 72 °C

Ars23SR 50 -GGTCCTCCAGTTAGTGTTACCCAAC-30 (Thao and Baumann, 2004) infB (Degenerate)

infB-1186-F 50 -ATYATGGGHCAYGTHGAYCAYGGHAARAC-30

3 min @ 95 °C; 34 cycles of 60s @ 94 °C, 60s @ 55 °C, 90s @ 72 °C; 10 min @ 72 °C

infB-1833-R 50 -TATCCGACGCCGAACTCCGRTTNCGCATNGC NCGNAYNCGNCC-30 (Hedegaard et al., 1999) infB (Arsenophonus)

A-infBF 50 -GATCCGGCCATACTCAAAAC-30

3 min @ 95 °C; 34 cycles of 60s @ 94 °C, 60s @ 53 °C, 90s @ 72 °C; 10 min @ 72 °C

A-infBR50 -GACCACGGCAAAACTTCATT-30 (This study) P45 (APSE)

APSE30.1 50 -ACGGCACTTAAACGCTATCC-30

2 min @ 94 °C; 36 cycles of 30s @ 94 °C, 50s @ 53 °C, 90s @ 72 °C; 5 min @ 72 °C

APSE31.1 50 -TGGGATGTGTATGGACGTTG-30 (Degnan and Moran, 2008) Histone H3 (insect)

H3aF 50 -ATGGCTCGTACCAAGCAGACVGC-30

3 min @ 94 °C; 40 cycles of 45s @ 94 °C, 30s @ 65 °C, 60s @ 72 °C; 6 min @ 72 °C

H3aR 50 -ATATCCTTRGGCATRATRGTGAC-30 (Colgan et al., 1998) M13 (plasmid)

M13F 50 -GTAAAACGACGGCCAGT-30 M13R 50 -GCGGATAACAATTTCACACAGG-30

arthropod symbionts and related members of the Enterobacteriaceae, whereas sequences in the P45 dataset were selected from GenBank solely by nucleotide similarity to those collected from the samples. The datasets also included infB and P45 sequences from an Arsenophonus-infected greenhouse whitefly (Trialeurodes vaporariorum; Taylor et al., unpublished). These were used to compare A. nasoniae with a separate species of Arsenophonus. The accession numbers for all database sequences used in the phylogenetic analysis are shown in Figs. 1 and 2. E. coli shows moderate levels of divergence from Arsenophonus and was used as the outgroup for the infB phylogeny. E. coli has previously been used to root 16S rDNA trees of Arsenophonus (Dale et al., 2006; Thao et al., 2000; Trowbridge et al., 2006). Phylogenetic trees for the APSE gene were visualized using midpoint rooting. For each gene the nucleotide DNA sequences were aligned using MUSCLE (Edgar, 2004), curated using Gblocks v0.91b (Talavera and Castresana, 2007) under the default settings, and output as a PHYLIP file. These alignments were manually confirmed using MacClade v.4.08, and the best model was found using Modeltest 3.7 (Posada and Crandall, 1998) in PAUP 4.0b10 (Swofford, 2003). Maximum likelihood phylogenies were constructed using the phylogeny.fr platform (Dereeper et al., 2008) using PHYML (Guindon and Gascuel, 2003; Guindon et al., 2005) with the optimum parameters as suggested by Modeltest. The infB phylogeny was performed with a GTR+C+I model using 427 nucleotide characters, with the following transition/transversion rates: AC:0.5310,

Used for sequencing clones

AG:2.7576, AT:0.7580, GC:0.4391, CT:2.7576, GT:1.0000. The parameters were set at 0.5612 for the proportion of invariable sites, and a gamma shape parameter (C) of 2.3964. The base frequencies were estimated from the data (A:0.2467, C:0.2326, G:0.2498, T:0.2709), and the final tree had a score of 2966. The maximum likelihood analysis for the APSE sequence was performed using 703 nucleotide bases under a GTR+C model with the following transition/transversion rates: AC:2.1017, AG:6.8375, AT:0.9115, GC:1.9309, CT:6.8375, GT:1.0000. A gamma shape parameter of 0.2319 was used, and with estimated base frequencies (A:0.2644, C:0.2406, G:0.2579, T:0.2371) the produced tree had a score of 2070. Node support for both trees was determined using 100 bootstrap replicates. 3. Results 3.1. Presence of A. nasoniae We identified infections of Arsenophonus in four of the 25 species of wasps tested; i.e., N. vitripennis, Spalangia endius, S. cameroni, Eupelmus vesicularis (Table 1). Despite being in the same superfamily (Chalcidoidea), these wasps are not closely related (Desjardins et al., 2007). Spalangia and Nasonia are in distantly related subfamilies in the Pteromalidae, and Eupelmus are in the family Eupelmidae. Infected individuals were collected from field populations in Canada, the United States, Romania, Turkey, and Peru. Infections

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Fig. 1. Maximum likelihood phylogeny of Arsenophonus nasoniae using the infB protein encoding gene. Numbers indicate bootstrap percentage at each node (out of 100 bootstraps). Only bootstrap values over 70 are displayed. NCBI accession numbers are shown in square brackets. For samples from this study, sampling location is reported.

Fig. 2. Midpoint rooted maximum likelihood phylogeny of the APSE P45 viral gene in a clade of insect symbionts. Host insect of each symbiont is identified in parenthesis. Numbers indicate bootstrap percentage at each node (out of 100 bootstraps). Only bootstrap values over 70 are displayed. NCBI accession numbers are shown in square brackets. For samples from this study, sampling location is reported.

were only detected in a portion of individuals, when multiple individuals were tested from the same population. Infections were detected in 47% (38 of 81) of female N. vitripennis field-collected at Lethbridge, AB. Infections were not detected in lab-cultured wasps, nor in any fly species. Sequences at the 16S rDNA locus from all wasp samples were identical over 453 nucleotide positions, and differed from the A. nasoniae type strain (accession number: AY264674.1) at one out of 453 sites. Because 16S rDNA is highly conserved, other Arsenophonus strains are nearly identical at this locus. The wasp samples also matched Arsenophonus from T. vaporariorum with 99% identity (identical at 372 out of 375 sites). The infB locus also showed high levels of similarity between samples. The infB sequences from three infected samples (N. vitripennis and E. vesicularis from Canada, S. cameroni from Russia) were identical to the type strain of A. nasoniae, whereas the other sample (E. vesicularis from Romania) differed at one silent site. Thus, all sequences obtained from parasitoid wasps clustered tightly together into a single clade (Fig. 1; 86% bootstrap support). The infB sequences from these wasps showed 95% identity to sequence from

the Arsenophonus from T. vaporariorum, differing at 21–22 out of 427 nucleotides. Sequence for the infB gene of S. endius samples was not obtained. 3.2. Prevalence of a phage sequence among individuals infected with A. nasoniae We identified phage sequence in E. vesicularis, N. vitripennis, and S. cameroni (Table 1). The fourth Arsenophonus-infected species, S. endius, exhibited the expected band but it was not confirmed by sequencing. Phage sequences were also identified from the type strain of A. nasoniae, as well as cultures that we isolated from N. vitripennis collected in the field. Not all Arsenophonus-infected individuals contained phage sequence (Table 1). All APSE-like sequences from the three wasp species were highly similar (kA: .008–.01 [4–5 out of 526 sites]; kS: 9–12% [15–19 out of 173 sites]), and clustered tightly together with high bootstrap support (99%) alongside the sequence from the A. nasoniae type strain (Fig. 2). The viral sequence found in the whitefly T. vaporariorum is allied with this clade (96% bootstrap support) (kA: .045–.052 [23–26

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out of 526 sites]; kS: .39–.44 [52–58 out of 173 sites]), and is 89% identical to the A. nasoniae type strain. All APSE-like sequences within these wasps are similar to APSE sequence found in H. defensa (kA: .035–.04 [18–21 out of 526 sites]; kS: .35–.4 [48–53 out of 173 sites]), with a maximum nucleotide divergence of 11%. 3.3. Screening for other potential APSE hosts It is unlikely that Sodalis is a major host for the APSE phage sequences we identified. Only the single S. cameroni sample from Russia tested positive for Sodalis using diagnostic 16S rRNA primers. All other APSE-positive wasp samples tested negative for this putative phage host. 4. Discussion Using primers for 16S rRNA and two variable protein-coding genes, we screened a total of 36 species of wasps and flies from 28 populations (16 countries) and detected infections of A. nasoniae in three parasitoid wasp species in addition to N. vitripennis. Recently, Duron et al. (2010) surveyed nine wasp and one fly species from 23 populations (two countries) and also found A. nasoniae infections in several species. Hence, this bacterium likely infects multiple hosts and should be considered a generalist, rather than a specialist of N. vitripennis. Although our screening method did not formally show that A. nasoniae resides within the insect body as a true symbiont, two lines of evidence support this conclusion. Firstly, wasp larvae purge gut contents prior to pupation (Tormos et al., 2009) and are unlikely to maintain contaminants across metamorphic stages. Although adult wasps could become contaminated through feeding on A. nasoniae infected hosts, the low prevalence of the bacterium in the field suggests that this is unlikely. Secondly, insect samples were washed prior to DNA extraction to reduce surface contaminants. Previous studies have found that A. nasoniae infects N. vitripennis at low frequencies (5–10% wasps; Balas et al., 1996). It is therefore perhaps surprising that we identified infected individuals in this study, especially since some of the populations we sampled had very small sample sizes. For example, some infected individuals are the sole representative of their population (i.e., E. vesicularis from Turkey, S. cameroni from Russia) and this suggests that A. nasoniae may occur in some species at much higher frequencies than previous reports. Infection frequencies in the field populations of N. vitripennis near Lethbridge, AB, were also high (47%). What accounts for this difference in infection frequencies amongst populations is unclear. Both different host backgrounds and bacterial strains may alter the phenotype presented. Although certain A. nasoniae strains cause male-killing, even upon transfer into novel hosts (Duron et al., 2010), not all A. nasoniae-infected wasps exhibit male-killing (Balas et al., 1996). The Lethbridge N. vitripennis population tested here has historically not exhibited significantly skewed sex-ratios (Floate et al., 1999). Phenotypic variation may alter selective pressures for the bacterium and affect the bacterium’s prevalence. Symbionts that require a selective pressure to remain in the host, i.e., those that alter host fitness or reproduction, are liable to be lost in the lab in the absence of environmental factors (Oliver et al., 2008). Supporting this, A. nasoniae was only found in field-collected individuals and not from laboratory maintained cultures. However, lab cultures are often established from few individuals, and the lack of infections may instead be a product of initial population sampling. When viewed as a primarily maternally inherited symbiont, it is not known how A. nasoniae persists in N. vitripennis in the longterm. No clear fitness benefit has been identified, although it has been proposed that male-killing may reduce inbreeding depression

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in Nasonia (Balas et al., 1996). However, it is not clear whether inbreeding depression is an important force in wild Nasonia (Luna and Hawkins, 2004). It is feasible that A. nasoniae is maintained within the community by horizontal transmission. This notion of a parasitic lifestyle is supported by three points. First, the broad host range of A. nasoniae suggests the bacterium spreads through several wasp and fly species in the environment. Second, the life history of A. nasoniae, both its transmission outside of the host as well as its ability to colonize novel hosts, suggest that the bacterium has a broader set of metabolic capabilities than most maternally transmitted insect symbionts. This is supported by the fact that the recently sequenced A. nasoniae genome contains diverse genes for carbohydrate metabolism (Darby et al., 2010). Finally, A. nasoniae can be horizontally transmitted under conditions of superparasitism to novel wasp species in the laboratory environment (Duron et al., 2010). This transfer is highly efficient, and the bacterium is subsequently maternally transmitted by the host with high fidelity. Killing males may be an incidental phenotype rather than a mechanism that is essential for the bacterium’s maintenance. However, it may have been necessary to invade the population initially. Of note is the ‘incremental gain hypothesis’, put forth by Balas et al. (1996). This hypothesis suggests a male-killing strain may invade a host population and supplant an existing benign strain if it provides a minor fitness benefit to the host. If this is the case, it may be difficult to detect fitness benefits for male-killing, because comparisons between the male-killing and non-male-killing strains are required. Regardless, future studies to determine the cost of infection to the host, and if A. nasoniae is maintained in the host in the absence of horizontal transmission, will be invaluable. The putative phage found in A. nasoniae may play an important role in the bacterium’s life history. It will be important to determine whether there are active phage particles infecting A. nasoniae, and if so, whether they are infectious and have a wide or narrow host range. It is also possible that we have identified sequences that are integrated into the bacterial genome and not associated with active phage. We also cannot exclude the possibility that the phage sequences we identified in some of the insects we screened are from a symbiont other than A. nasoniae. However, two lines of evidence suggest otherwise. First, there was a very strong correlation between A. nasoniae infection and P45 sequence. Although SuPER PCR identified a Sodalis-like sequence in S. cameroni, this was not found in any other APSE-positive samples. Second, and more importantly, we were able to obtain P45 sequence from A. nasoniae cultures. Although some phages have broad host ranges (Jensen et al., 1998), most are restricted to closely-related hosts (Welkos et al., 1974); e.g., two phages related to APSE infect single hosts within the Enterobacteriaceae (Clark et al., 2001; Vander Byl and Kropinski, 2000). It is not clear why P45 was not amplified from all A. nasoniae samples. This may be caused by either sequence divergence at the primer binding locations, or by a secondary loss of the phage (Degnan and Moran, 2008). The phage sequences identified in this study are highly similar to those found in two other defensive symbionts – a strain of H. defensa that protects the pea aphid, Acyrthosiphon pisum, against parasitic wasps (Oliver et al., 2009), and a strain of Arsenophonus that infects the psyllid Glycaspis brimblecombei, and whose prevalence is correlated with parasitism pressure (Hansen et al., 2007). This suggests the phage sequence in A. nasoniae may have potential ecological implications. It is important to determine if the phage locus in A. nasoniae represents a functional phage and expresses proteins within the host. Because toxins produced by APSE in the pea aphid prevent wasps from developing in the host (Oliver et al., 2009), it is tantalizing to hypothesize that the mechanisms for phage-mediated defence and male-killing may be similar.

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No fitness benefit has been found for A. nasoniae infection in N. vitripennis, but the bacterium infects multiple wasp species in the environment. This broad host range suggests horizontal transmission may play an integral role in bacterial maintenance. The importance of horizontal transmission remains to be directly tested, and the bacterium may have significant impacts on novel hosts in the environment – particularly given the lysogenic phage potentially present. Perplexingly, the Son-killer bacterium is defined by its male-killing phenotype, but the importance of this reproductive manipulation for its maintenance remains completely unknown. Acknowledgments For providing insect samples, we are grateful to A. Oliva, A. Thompson, S. Barker, C. Covacin, G. Boivin, R. Bourchier, B. Broadbent, H. Cárcamo, D. Colwell, J. Cossentine, R. De Clerck-Floate, K. Deglow, G. Gibson, M. Goettel, T. Lowery, P. Mason, O. Olfert, J. Soroka, L. Wambold, J. Whistlecraft, H. Philips, S. Gutierrez, A. Schaaf, C. Roy, B. McCron, B. Hall, D. Kimball, B. Cade, S. Walker, M-P. Mignault, M. Reid, H. Skovgård, J. Fourrier, P.S. Castillo Carrillo, W. Blanckenhorn, S. Penn, T. McKay, C.J. Geden, D. Taylor, D. Bragg, J. Hogsette, R. Moon, M. Hoffman, B. Pawson, B.H. King, L. Fuzu, J. Werren, and P. Kaufman. The comments of two anonymous reviewers greatly improved the manuscript. This research was funded by NSERC and Agriculture and Agri-Food Canada. Steve Perlman acknowledges support from the Canadian Institute for Advanced Research. References Andreadis, T.G., 1985. Epizootiology of Amblyospora stimuli (Microsporidiida: Amblyosporidae) infections in field populations of a univoltine mosquito, Aedes stimulans (Diptera: Culicidae), inhabiting a temporary vernal pool. J. Invertebr. Pathol. 74, 198–205. Balas, M.T., Lee, M.H., Werren, J.H., 1996. Distribution and fitness effects of the sonkiller bacterium in Nasonia. Evol. Ecol. 10, 593–607. Baumann, P., Lai, C.Y., Baumann, L., Rouhbakhsh, D., Moran, N.A., Clark, M.A., 1995. Mutualistic associations of aphids and prokaryotes – biology of the genus Buchnera. Appl. Environ. Microbiol. 61, 1–7. Clark, A.J., Inwood, W., Cloutier, T., Dhillon, T.S., 2001. Nucleotide sequence of coliphage HK620 and the evolution of lambdoid phages. J. Mol. Biol. 311, 657– 679. Colgan, D.J., McLauchlan, A., Wilson, G.D.F., Livingston, S.P., Edgecombe, G.D., Macaranas, J., Cassis, G., Gray, M.R., 1998. Histone H3 and U2 snRNA DNA sequences and arthropod molecular evolution. Aust. J. Zool. 46, 419–437. Dale, C., Beeton, M., Harbison, C., Jones, T., Pontes, M., 2006. Isolation, pure culture, and characterization of ‘‘Candidatus Arsenophonus arthropodicus’’, an intracellular secondary endosymbiont from the hippoboscid louse fly Pseudolynchia canariensis. Appl. Environ. Microbiol. 72, 2997–3004. Darby, A., Choi, J., Wilkes, T., Hughes, M., Werren, J.H., Hurst, J., Colbourne, J., 2010. Characteristics of the genome of Arsenophonus nasoniae, son-killer bacterium of the wasp Nasonia. Insect Mol. Biol. 19, 75–89. Degnan, P.H., Moran, N.A., 2008. Evolutionary genetics of a defensive facultative symbiont of insects: exchange of toxin-encoding bacteriophage. Mol. Ecol. 17, 916–929. Dereeper, A., Guignon, V., Blanc, G., Audic, S., Buffet, S., Chevenet, F., Dufayard, J.F., Guindon, S., Lefort, V., Lescot, M., Claverie, J.M., Gascuel, O., 2008. Phylogeny.fr: robust phylogenetic analysis for the non-specialist. Nucleic Acids Res. 36, W465–W469. Desjardins, C.A., Regier, J.C., Mitter, C., 2007. Phylogeny of pteromalid parasitic wasps (Hymenoptera: Pteromalidae): initial evidence from four protein-coding nuclear genes. Mol. Phylogenet. Evol. 45, 454–469. Duron, O., Bouchon, D., Boutin, S., Bellamy, L., Zhou, L., Engelstadter, J., Hurst, G.D., 2008. The diversity of reproductive parasites among arthropods: Wolbachia do not walk alone. BMC Biol. 6, 27. Duron, O., Wilkes, T.E., Hurst, G.D.D., 2010. Interspecific transmission of a malekilling bacterium on an ecological timescale. Ecol. Lett. 13, 1139–1148. Edgar, R.C., 2004. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res. 32, 1792–1797. Ferree, P.M., Avery, A., Azpurua, J., Wilkes, T., Werren, J.H., 2008. A bacterium targets maternally inherited centrosomes to kill males in Nasonia. Curr. Biol. 18, 1409– 1414. Floate, K.D., 2002. Production of filth fly parasitoids (Hymenoptera: Pteromalidae) on fresh and on freeze-killed and stored house fly pupae. Biocontrol Sci. Technol. 12, 595–603. Floate, K.D., Khan, B., Gibson, G., 1999. Hymenopterous parasitoids of filth fly (Diptera: Muscidae) pupae in cattle feedlots. Can. Entomol. 131, 347–362.

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