Archives of Virology

, Volume 162, Issue 2, pp 449–456 | Cite as

Discovery of herpesviruses in Canadian wildlife

  • Chimoné S. Dalton
  • Karen van de Rakt
  • Åsa Fahlman
  • Kathreen Ruckstuhl
  • Peter Neuhaus
  • Richard Popko
  • Susan Kutz
  • Frank van der Meer
Original Article


Herpesviruses (HVs) have a wide range of hosts in the animal kingdom. The result of infection with HVs can vary from asymptomatic to fatal diseases depending on subtype, strain, and host. To date, little is known about HVs naturally circulating in wildlife species and the impact of these viruses on other species. In our study, we used genetic and comparative approaches to increase our understanding of circulating HVs in Canadian wildlife. Using nested polymerase chain reaction targeting a conserved region of the HV DNA polymerase gene, we analyzed material derived from wildlife of western and northern Canada collected between February 2009 and Sept 2014. For classification of new virus sequences, we compared our viral sequences with published sequences in GenBank to identify conserved residues and motifs that are unique to each subfamily, alongside phylogenetic analysis. All alphaherpesviruses shared a conserved tryptophan (W856) and tyrosine (Y880), betaherpesviruses all shared a serine (S836), and gammaherpesviruses had a conserved glutamic acid (E835). Most of our wildlife HV sequences grouped together with HVs from taxonomically related host species. From Martes americana, we detected previously uncharacterized alpha- and beta-herpesviruses.


Bighorn Sheep Rangifer Tarandus Canadian Wildlife Bayesian Phylogenetic Tree Rangifer Tarandus Tarandus 
These keywords were added by machine and not by the authors. This process is experimental and the keywords may be updated as the learning algorithm improves.


Herpesviridae is a family of enveloped, double-stranded DNA viruses that emerged roughly 400 million years ago [1]. Herpesviruses (HVs) closely co-evolved with their natural human and animal hosts and can be classified into three subfamilies: Alpha-, Beta-, and Gammaherpesvirinae [2, 3, 4]. This classification is based on phylogeny, pathogenesis, and the organ or tissue in which these viruses establish latency: neural ganglia, secretory glands, and lymphatic tissues, respectively [2, 4, 5, 6]. Transmission of HV may occur vertically or horizontally in a population where susceptible animals become infected through direct contact with mucosal surfaces of infected individuals [7]. In the event of cross-species HV transmission, disease is often characterized by lesions, fever, weight loss, spontaneous abortions, and death [8, 9].

In northern Alberta, Canada, serological tests have confirmed the presence of alphaherpesviruses in woodland caribou (Rangifer tarandus caribou) [10]. Koi herpesvirus (KHV) was detected in wild carp (Cyprinus carpio) of Manitoba and Ontario through PCR and sequencing [11]. Phocid herpesvirus 1 (PhHV-1), identified through microscopic evidence of inclusion bodies, caused morbidity and mortality in neonatal and weaning harbor seal (Phoca vitulina) pups of British Columbia, Canada and Washington, USA [12]. In Calgary, Alberta, Canada, Washington, USA, and Idaho, USA, numerous great horned owls (Bubo virginianus), peregrine falcons (Falco peregrinus), and gyrfalcons (Falco rusticolus) have died from infection with columbid herpesvirus 1 originating from pigeons (Columba livia domestica) diagnosed though sequencing of HV genetic material [13, 14]. Awareness of circulating viruses could lead to targeted post-mortem examinations, thereby improving wildlife disease surveillance [15].

The HV DNA polymerase gene can be detected by PCR, using published degenerate primers [16]. The HV DNA polymerase gene, which encodes a B family polymerase, is common to all HVs regardless of subfamily or strain. Although DNA polymerase genes of HVs are not identical, they are structurally related and contain highly conserved B (region III) and C (region I) motifs, which are targeted by this PCR assay [17]. Motif B comprises the sequence KXXXNSXYGXXG and is believed to be a part of the polymerase active site involved in dNTP coordination and DNA synthesis [17, 18]. Motif C is composed of the sequence DTDS, which, together with motif A, is critical for cation coordination when the enzyme is active [17]. Mutations in these known conserved motifs lead to severe impairment of the viral polymerase enzyme activity [19].

Our study analysed the diversity of circulating HVs in selected and opportunistically obtained samples collected between 2010 and 2014 from Canadian native wildlife. We used phylogenetic approaches to compare HVs and found two previously uncharacterized marten HV sequences among our samples.

Materials and methods

Sample collection

Convenience sampling methods were used to collect tissue and blood samples from various Canadian wildlife animals between February 2009 and September 2014. Animals presented for necropsy to the University of Calgary originated from research, trappers, hunting, roadkill events, euthanasia, or wildlife management activities. Tissue samples were taken from spleen, mesenteric lymph nodes, liver, kidney, tonsil and lung during post-mortem examinations. When possible, fresh whole blood was collected into serum and/or EDTA tubes.

Sample preparation and DNA extraction

Animal tissue pieces of approximately 0.3 g were finely minced with a scalpel blade and placed in a 1.5-mL microcentrifuge tube. DNA was extracted using an E.Z.N.A.® Tissue DNA Kit (Omega Bio-Tek Inc., Norcross, GA, USA) following the manufacturer’s protocol. Prior to extraction, tissue pieces were incubated overnight at 55 °C with TL buffer to achieve optimal tissue digestion. DNA was eluted twice from a HiBind DNA mini column, each time using 25 µL of 10 mM Tris HCl, pH 8.5, at 70 °C.

Serum was collected from coagulated whole blood by centrifugation for 10 minutes at 1,000×g. Peripheral blood mononuclear cells (PBMC) were isolated from EDTA blood by density gradient centrifugation on Ficoll-Paque® PLUS (GE Healthcare Bio-Sciences AB, Uppsala, Sweden). Viral DNA was extracted using an E.Z.N.A.® Blood DNA Kit (Omega Bio-Tek Inc., Norcross, GA, USA).

Polymerase chain reaction conditions

Nested degenerate primers were used to amplify a highly conserved fragment between motifs B and C of the DNA-directed DNA polymerase gene common to all HVs [16]. These primers allow rapid amplification of the polymerase gene of all known HV subfamilies. For primary amplification, we used 75-150 ng of template DNA, 1.25 units of AccuTaq™ LA DNA Polymerase (Sigma-Aldrich Co. LLC, St. Louis, MO, USA), 3.75 pmol of each nucleotide, and 5 pmol each of primers DFA, KG1, and ILK in a total reaction volume of 25 µL. Cycling conditions were as follows: initial denaturation of 5 min at 95 °C followed by 40 cycles of 95 °C for 30 s, 46 °C for 60 s, and 72 °C for 90 s, with a final extension step of 5 min at 72 °C. The second round of the nested protocol used 1.5 µL of the first-round PCR product as template with 5 pmol of primers IYG and TGV. Thermocycling conditions were as described above, but with 30 seconds of annealing time followed by 60 seconds of extension at each cycle.

Gel extraction and sequencing

PCR products were separated using gel electrophoresis on a 1% agarose gel, and bands were extracted using an E.Z.N.A.® Gel Extraction Kit (Omega Bio-Tek Inc., Norcross, GA, USA), following the manufacturer’s protocol. DNA was eluted from the minicolumn using 30 µL of elution buffer heated to 70 °C. DNA samples were premixed with IYG primer and sequenced (Eurofins MWG Operon LLC, Huntsville, AL, USA). Weak amplification bands were purified from the gel, and the DNA was ligated into PGEM-T Easy Vector (Promega Corp. Madison, WI, USA), followed by transformation using One Shot® TOP10 E. coli (Invitrogen Corp., Carlsbad, CA, USA). LB agar plates with 1% ampicillin and spread with 1.6 mg X-GAL were used for blue-white colony screening. Five white colonies were grown in LB with 1% ampicillin. Plasmids were recovered using an E.Z.N.A.® Plasmid Mini Kit (Omega Bio-Tek Inc., Norcross, GA, USA) following the manufacturer’s protocol and sequenced using T7 and SP6 primers (Eurofins MWG Operon LLC, Huntsville, AL, USA).

Phylogenetic analysis

Chromatograms of sequences were visually inspected for quality. Poor-quality nucleotide sequences and short sequences were discarded from the analysis. Fragments generated from T7 and SP6 primers were used to construct a consensus DNA fragment for each replicate of each sample. Primer and vector sequences were trimmed prior to alignment using Geneious v8.1.3 [20]. Only amino acid sequences that covered the full range between motif B and C on the DNA polymerase gene were included in further analysis. These viral sequences were searched against published sequences in GenBank using the protein Basic Local Alignment Search Tool (BLASTp) [21]. Additional known DNA polymerase sequences of HVs were obtained from GenBank and included in amino acid alignments using the multiple sequence comparison by log-expectation (MUSCLE) algorithm [22]. Since this region of the viral DNA polymerase gene is commonly used to detect HVs, amino acid sequences of wildlife and published HV sequences were aligned to examine conserved residues and motifs characteristic of each subfamily. Phylogenetic trees were constructed using the Randomized Axelerated Maximum Likelihood (RAxML) tree builder with 100 bootstrap replicates [23], and also using the MrBayes Bayesian inference plugin available in Geneious with a burn-in of 1000. Tree files were exported and visually labelled for clarity in FigTree v1.4.2 [24].


A fragment of the HV DNA-dependent DNA polymerase gene was successfully amplified by nested PCR from wildlife tissue and blood samples from different wildlife species. High-quality nucleotide sequences were obtained from 21 animals of six different species, and these are summarised in Table 1 along with their most similar sequence found using BLASTp. Tissue samples from other Porcupine caribou herd members (Rangifer tarandus granti), fox (Vulpes vulpes), wolf (Canis lupus), cougar (Puma concolor), black bear (Ursus americanus), grizzly bear (Ursus arctos horribilis), and raccoon (Procyon lotor) were negative using the described PCR methodology. Overall, the use of spleen tissue resulted in the largest number of successful sequencing attempts when compared to other tissue types. Fifty-one sequences were included in the MUSCLE amino acid alignment and RAxML phylogenetic tree (Fig. 1). These same sequences were included in Bayesian phylogenetic analysis to strengthen our interpretations. Similar topologies were seen for both phylogenetic trees, with greater branching support in the Bayesian tree. Three major viral groups formed in both trees, corresponding to the HV subfamilies Alpha-, Beta-, and Gammaherpesvirinae. However, FHV-1 did not follow this topology in the RAxML analysis.
Table 1

A summary of herpesvirus DNA-dependent DNA polymerase gene amino acid sequences from this study and the most closely related virus sequences in the GenBank database

Host organism


GenBank accession number

Most similar BLASTpa match (accession no., % identity)

Bighorn sheep (Ovis canadensis)

5, 13

KX062141, KX062142

Ruminant rhadinovirus 2, bighorn sheep (AAO88175, 98%)

Bison (Bison bison)



American bison gammaherpesvirus (AAL29891, 100%)

Caribou, Porcupine herd (PCH, Rangifer tarandus granti)



Reindeer gammaherpesvirus (AFV98876, 100%)

21, 24

KX062138, KX062139

Reindeer gammaherpesvirus (AF11CanadaV98876, 98%)

Coyote (Canis latrans)

12, 27

KX062143, KX062144

Canid herpesvirus 1 (AAC55646, 100%)

Elk (Cervus canadensis)

11, 13

KX062145, KX062146

Ruminant rhadinovirus 2, elk (AAO88180, 95%)



Ruminant rhadinovirus 2, elk (AAO88180, 100%)



Ruminant rhadinovirus 2, elk (AAO88180, 97%)

Marten (Martes americana)

8, 32

KX062128, KX062130

Mustelid herpesvirus 1 (AAL55728, 93%)

33, 41, 46

KX062131, KX062132, KX062133

Felid herpesvirus 1 (AAC55649, 72%)

11, 49, 52, 65

KX062129, KX062134, KX062135, KX062136

Aotine herpesvirus 1(AAC55643, 52%)

a GenBank protein Basic Local Alignment Search Tool [21]

Fig. 1

RAxML tree of a MUSCLE alignment of 51 herpesvirus DNA polymerase amino acid sequences. Twenty-one viral sequences from wildlife from this study are shown in blue, and accession numbers are shown in parentheses. PCH represents caribou from the Porcupine herd. Maximum-likelihood analysis was carried out in Geneious 8.1 using the RAxML plugin, with the scale bar indicating the number of amino acid substitutions per site. Branching support is shown as bootstrap percentages for 100 bootstrap replicates (color figure online)

Bison (Bison bison) sample 15 was positive for American bison gammaherpesvirus (AAL29891, 100%), and our four elk (Cervus canadensis) samples were infected with variant strains of ruminant rhadinovirus 2 of elk (AAO88180, 95%-100%) as summarized in Table 1.

Two coyote (Canis latrans) samples from Alberta, Canada, shared 100% nucleic acid and amino acid identity with canid herpesvirus 1 (CHV-1) (accession number AAC55646) isolated from domestic Labrador retriever (Canis lupus familiaris) puppies. CHV-1 is an alphaherpesvirus known to infect domestic and wild canines. Coyote sequences clustered within the alphaherpesvirus subfamily with CHV-1 in phylogenetic analysis (Figs. 1 and 2).
Fig. 2

MrBayes Bayesian inference phylogenetic tree of a MUSCLE alignment of 51 herpesvirus DNA polymerase amino acid sequences. Twenty-one viral sequences from wildlife from this study are shown in blue, and accession numbers are shown in parentheses. PCH represents caribou from the Porcupine herd. Bayesian analysis was carried out in Geneious 8.1 using the MrBayes plugin, with the scale bar indicating the number of amino acid substitutions per site. A burn-in of 1,000 was used, with branching support shown as posterior probability percentages (color figure online)

Marten (Martes americana) samples were collected from trappers in the Sahtú Settlement region, Northwest Territories, Canada. Amino acid and nucleic acid sequences from martens 8 and 32 were identical and shared 93% identity with mustelid herpesvirus 1 (AAL55728), a gammaherpesvirus found in sea otters (Enhydra lutris) and other members of the family Mustelidae such as badgers (Meles meles) and pine martens (Martes martes) of Europe.

Bighorn sheep (Ovis canadensis) sample 5 and sample 13, native to Sheep River Provincial Park, Alberta, Canada, shared 98% amino acid sequence identity with the gammaherpesvirus ruminant rhadinovirus 2 (AAO88175) from free-range bighorn sheep in Washington, USA [25].

Marten samples 33, 41, and 46 shared 72% identity with the alphaherpesvirus felid herpesvirus 1 (AAC55649), which infects domestic cats. The amino acid sequence identified from marten 46 differed from those from martens 33 and 41 by one residue at position 38; the nucleotide sequence CTT in marten 46 was changed to CCT, thereby encoding a proline instead of a leucine at this position. Martens 33, 41, and 46 composed a single phyletic group within the subfamily Betaherpesvirinae (Figs. 1 and 2).

Marten samples 11, 49, 52, and 65 were infected with an uncharacterized betaherpesvirus. Sequences from these marten sequences are identical in amino acid sequence and have no close relatives in GenBank (Table 1). These martens aligned in a single phyletic group within the subfamily Betaherpesvirinae; however, other sequences in this subfamily are distant (Figs. 1 and 2).

Spleen samples from animals 15, 21, and 24 of the Porcupine caribou herd (PCH) native to northwestern Canada, produced amino acid sequences that were 98-100% identical to those of reindeer gammaherpesvirus (AFV98876) characterized in Norway from caribou (Rangifer tarandus tarandus) [26]. Sample PCH 21 differed from the Norwegian virus by an amino acid mutation from an isoleucine to a valine, sharing 98.3% identity. PCH sample 24 was more variable in the terminal region of the fragment, differing from those from other herd members and reindeer gammaherpesvirus.

Starting at the first residue after the TGV primer corresponding to position 824 of the polymerase, all sequences shared a proline (P829), ?cysteine? (C830), alanine (A834), threonine (T838), glycine (G841), serine (S842), methionine (M844), and leucine (L845). Variation in the amino acid sequence occurs more frequently in the terminal end of the fragment between motif B (region III) approaching motif C (region I) of the DNA polymerase polypeptide.

The sequences from martens 11, 33, 41, 46, 49, 52, and 65 were aligned with most similar published sequences in GenBank using MUSCLE to highlight amino acid sequence differences (Fig. 3). Published alphaherpesvirus sequences share a conserved aromatic hydrophobic tryptophan (W856) residue and a tyrosine (Y880) at a position five residues from the end of the sequence. The previously undescribed sequences from martens 33, 41, and 46 in this region are most similar to those of alphaherpesviruses, since they align with the alphaherpesvirus sequences in maximum-likelihood and Bayesian phylogenetic trees and share subfamily-specific conserved residues.
Fig. 3

Alpha- and betaherpesvirus-like marten amino acid sequences aligned with the most closely related published sequences using MUSCLE

Viral sequences from the spleens of martens 11, 49, 52, and 65 are most similar to betaherpesviruses, since they share the subfamily-specific conserved serine (S836) residue at position 13. These beta-like marten sequences also cluster with other betaherpesvirus sequences in maximum-likelihood and Bayesian phylogenetic trees, although the relationship is distant in both.

Published gammaherpesvirus sequences all share a glutamic acid (E835) residue at position 12. Our previously undescribed viral sequences identified in martens 8 and 32, bighorn sheep 5 and 13, elk 11, 13 and 49, and PCH 21 and 24 belong to the subfamily Gammaherpesvirinae based on conserved residues and clustering in both phylogenetic trees.


The Porcupine caribou herd spans Alaska, USA, and the northwest Yukon, Canada [27]. Evidence from this study suggests that reindeer gammaherpesvirus and at least one other closely related virus is circulating in the Porcupine caribou herd. Our Canadian caribou HV sequences share an ancestor with the Norway strain, which is not surprising, as North American caribou originally came from Eurasia [28].

Coyote samples in our study were infected with CHV-1, which is known to cause morbidity and mortality in wild and domestic Canidae pups. Coyotes are small canids native to North America that were historically confined to open plains and arid regions. With the expansion of urban developments, it is possible that CHV-1 was transmitted from domestic dogs to wild coyotes, and as such, CHV-1 was carried into the wild-coyote population. Wild coyotes are known to interact with wolves and foxes, both of which also have large territorial ranges [29]. Since coyotes, wolves, and foxes are susceptible to CHV-1, a single spillover event from domestic dogs could propagate disease in many animal populations.

As expected, our bighorn sheep, bison, and elk samples were infected with HVs characterized in previous studies from members of the same species. It is possible that related strains of ruminant rhadinovirus 2 are endemic in bison and elk, while American bison gammaherpesvirus is endemic in bison.

Martens are mid-sized carnivorous mammals that are distributed across forests of Canada and Alaska [30]. They are integrated in various habitats in urban and rural communities and hunt on the ground or in trees. These animals are solitary and territorial, with some dispersing over 80 km [30]. We identified different marten herpesviruses that segregated into all three HV subfamilies. Marten gammaherpesvirus sequences are closely related to mustelid HVs, which is as expected, since martens are of the Mustelidae animal family. In some cases, HV in martens are distantly related to other HVs in the alpha and beta subfamilies, likely because such viral sequences have not yet been characterized in closely related animals.

The fragment between the B and C motif contains several amino acid residues that are conserved within particular HV subfamilies, as confirmed in this study. It has been shown in studies with human herpesvirus 1 that a glycine-to-serine mutation at position 841 in the polymerase results in altered drug sensitivity [31]. It is possible that mutations in other conserved residues common to all HVs could also directly impact the function of the polymerase or the virus itself. The influence of subfamily-specific conserved residues identified in this study (alpha: W856, Y880; beta: S836; gamma: E835) on the function of the polymerase is also unknown but warrants investigation. The terminal region approaching motif C shows greater variability, suggesting that this region may not be critical for polymerase function. Future studies could determine the role of this region in infectivity or pathogenesis.

Convenience sampling allowed us to access tissues from numerous wildlife species, including more-scarce tissues from cougar, wolf, fox, racoon, and bear. However, many of these tissues tested negative when using the described methods. The high frequency of negative PCR results could be due to the handling, storage, or age of tissues prior to testing for HV. Plumb et al. showed in 1973 that the infectivity and detectability of channel catfish herpesvirus (CCV) in recently deceased channel catfish (Ictalurus punctatus) was greatly influenced by the storage temperature and duration of storage [32]. After 48 hours at 22 °C, CCV in dead fish was no longer infectious in cell culture. CCV could be detected up to 14 days when dead fish were stored on ice, up to 162 days when stored at –20 °C, and up to 210 days when the fish were frozen at –80 °C. Some of our wildlife samples were sourced from carcasses that were found dead, or from remote regions where access to a freezer was not always possible. The ability to detect HVs using PCR could be affected by the time of death and status of animal and viral degradation. Biopsies of available tissues were sometimes taken in non-sterile, wilderness settings, potentially introducing PCR inhibitors that might have contributed to the negative PCR results. Uniform sampling from different tissues of different animals, followed by immediate PCR analysis or proper tissue storage may reveal HVs present in species that tested negative in our study.

It is important that we understand the relationships between viruses and animals in a wild or domestic setting so that we can examine the overall health status of a population. In this study, we determined the nucleotide and predicted amino acid sequences of previously undescribed HVs in Canadian wildlife. Observing the conserved motifs of the HV DNA polymerase gene and additional conserved subfamily-specific residues allowed us to determine the relationship of these unknown HVs to known viruses. Amplifying larger DNA fragments of the HV genome will allow stronger evolutionary comparisons of HV genes and may give insights regarding the relatedness of host species harboring these co-evolving viruses. Finally, samples from other wild species and uniform sampling are needed for phylogenetic resolution when comparing previously undescribed viruses to known viruses.



We would like to thank the members of the Canadian Cooperative Wildlife Health Centre and the pathology department at the University of Calgary Spy Hill Campus for their help and expertise in animal sample collection during necropsy. Caribou samples were collected through CARMA project; some marten samples were collected by the youth of the Sahtú region as part of an NSERC PromoScience outreach program. Finally, Alasdair Veitch, Ale Massolo, and Cynthia Kashivakura participated largely in this project.

Compliance with ethical standards

Conflict of interest

The authors declare that they have no conflicts of interest.


This study was funded by University Research Grants Committee (URGC) Seed Grant Program, University of Calgary, Alberta, Canada.


  1. 1.
    McGeoch DJ, Gatherer D (2005) Integrating reptilian herpesviruses into the family Herpesviridae. J Virol 79:725–731CrossRefPubMedPubMedCentralGoogle Scholar
  2. 2.
    McGeoch DJ, Rixon FJ, Davison AJ (2006) Topics in herpesvirus genomics and evolution. Virus Res 117:90–104CrossRefPubMedGoogle Scholar
  3. 3.
    Wang N, Baldi PF, Gaut BS (2007) Phylogenetic analysis, genome evolution and the rate of gene gain in the Herpesviridae. Mol Phylogenet Evol 43(3):1066–1075CrossRefPubMedGoogle Scholar
  4. 4.
    Blake N (2010) Immune evasion by gammaherpesvirus genome maintenance proteins. J Gen Virol 91(4):829–846CrossRefPubMedGoogle Scholar
  5. 5.
    McGeoch DJ, Cook S, Dolan A, Jamieson FE, Telford EA (1995) Molecular phylogeny and evolutionary timescale for the family of mammalian herpesviruses. J Mol Biol 247:443–458CrossRefPubMedGoogle Scholar
  6. 6.
    McGeoch DJ, Dolan A, Ralph AC (2000) Toward a comprehensive phylogeny for mammalian and avian herpesviruses. J Virol 74:10401–10406CrossRefPubMedPubMedCentralGoogle Scholar
  7. 7.
    Wald A, Corey L (2007) Persistence in the population: epidemiology, transmission. In: Arvin A et al (ed) Human herpesviruses: biology, therapy, and immunoprophylaxis. Cambridge University Press, Cambridge, ch 36Google Scholar
  8. 8.
    Lankester F, Lugelo A, Kazwala R, Keyyu J, Cleaveland S, Yoder J (2015) The economic impact of malignant catarrhal fever on pastoralist livelihoods. PLoS One 10(1):e0116059CrossRefPubMedPubMedCentralGoogle Scholar
  9. 9.
    Mlilo D, Mhlanga M, Mwembe R, Sisito G, Moyo B, Sibanda B (2015) The epidemiology of malignant catarrhal fever (MCF) and contribution to cattle losses in farms around Rhodes Matopos National Park, Zimbabwe. Trop Anim Health Prod 47(5):989–994CrossRefPubMedGoogle Scholar
  10. 10.
    Tessaro SV, Deregt D, Dzus E, Rohner C, Smith K, Gaboury T (2005) Herpesvirus infection in woodland caribou in Alberta, Canada. J Wild Dis 41(4):803–805CrossRefGoogle Scholar
  11. 11.
    Garver KA, Al-Hussinee L, Hawley LM, Schroeder T, Edes S, LePage V et al (2010) Mass mortality associated with koi herpesvirus in wild common carp in Canada. J Wild Dis 46(4):1242–1251CrossRefGoogle Scholar
  12. 12.
    Himworth CG, Haulena M, Lambourn DM, Gaydos JK, Huggins J, Calambokidis J et al (2010) Pathology and epidemiology of Phocid herpesvirus-1 in wild and rehabilitating harbor seals (Phoca vitulina richardsi) in the northeastern Pacific. J Wild Dis 46(3):1046–1051CrossRefGoogle Scholar
  13. 13.
    Gailbreath K, Oaks L (2008) Herpesviral inclusion body disease in owls and falcons is caused by the pigeon herpesvirus (Columbid herpesvirus 1). J Wildl Dis 44:427–433CrossRefPubMedGoogle Scholar
  14. 14.
    Rose N, Warren AL, Whiteside D, Bidulka J, Robinson JH, Illanes O, Brookfield C (2012) Columbid herpesvirus-1 mortality in great horned owls (Bubo virginianus) from Calgary, Alberta. Can Vet J 53:265–268PubMedPubMedCentralGoogle Scholar
  15. 15.
    Brown M, Moore L, McMahon B, Powell D, LaBute M, Hyman JM et al (2015) Constructing rigorous and broad biosurveillance networks for detecting emerging zoonotic outbreaks. PLoS One 10(5):e0124037CrossRefPubMedPubMedCentralGoogle Scholar
  16. 16.
    Vandevanter DR, Warrener P, Bennett L, Schultz ER, Coulter S, Garber RL et al (1996) Detection and analysis of diverse herpesviral species by consensus primer PCR. J Clin Microbiol 34(7):1666–1671PubMedPubMedCentralGoogle Scholar
  17. 17.
    Bennett N, Götte M (2013) Utility of bacteriophage RB69 polymerase gp43 as a surrogate enzyme for herpesvirus orthologs. Viruses 5:54–86CrossRefPubMedPubMedCentralGoogle Scholar
  18. 18.
    Marchler-Bauer A, Derbyshire MK, Gonzales NR, Lu S, Chitsaz F, Geer LY et al (2015) CDD: NCBI’s conserved domain database. Nucleic Acids Res 28(43):D222. doi: 10.1093/nar/gku1221 CrossRefGoogle Scholar
  19. 19.
    Ye L-B, Huang E (1993) In vitro expression of the human cytomegalovirus DNApolymerase gene: Effects of sequence alterations on enzyme activity. J. Virol 67:6339–6347PubMedPubMedCentralGoogle Scholar
  20. 20.
    Kearse M, Moir R, Wilson A, Stones-Havas S, Cheung M, Sturrock S et al (2012) Geneious Basic: an integrated and extendable desktop software platform for the organization and analysis of sequence data. Bioinf 28(12):1647–1649CrossRefGoogle Scholar
  21. 21.
    NCBI (2013) National Center for Biotechnology Information BLAST home.
  22. 22.
    Edgar RC (2004) MUSCLE: multiple sequence alignment with high accuracy and high 180 throughput. Nucleic Acids Res 32(5):1792–1797CrossRefPubMedPubMedCentralGoogle Scholar
  23. 23.
    Stamatakis A (2014) RAxML Version 8: A tool for Phylogenetic Analysis and Post-Analysis of Large Phylogenies. Bioinf 30(9):1312–1313CrossRefGoogle Scholar
  24. 24.
    Morariu VI, Srinivasan BV, Raykar VC, Duraiswami R, Davis LS (2008) Automatic online tuning for fast Gaussian summation. Adv Neur Inf Proc Sys (NIPS).
  25. 25.
    Li H, Gailbreath K, Flach EJ, Taus NS, Cooley J, Keller J et al (2005) A novel subgroup of rhadinoviruses in ruminants. J Gen Virol 86:3021–3026CrossRefPubMedGoogle Scholar
  26. 26.
    das Neves CG, Ihlebaek HM, Skjerve E, Hemmingsen W, Li H, Tryland M (2013) Gammaherpesvirus infection in semi-domesticated reindeer (Rangifer tarandus tarandus): a cross-sectional, serologic study in northern Norway. J Wild Dis 49(2):261–269Google Scholar
  27. 27.
    COSEWIC (2011) Designatable units for caribou (Rangifer tarandus) in Canada. Committee on the Status of Endangered Wildlife in Canada: Ottawa, p 88.
  28. 28.
    Yannic G, Pellissier L, Ortego J, Lecomte N, Couturier S, Cuyler C (2014) Genetic diversity in caribou linked to past and future climate change. Nat Clim Change 4:132–137CrossRefGoogle Scholar
  29. 29.
    Levi T, Wilmers CC (2012) Wolves-coyotes-foxes: a cascade among carnivores. Ecology 93(4):921–929CrossRefPubMedGoogle Scholar
  30. 30.
    Broquet T, Johnson CA, Petit E, Thompson I, Burel F, Fryxell JM (2006) Dispersal and genetic structure in the American marten, Martes americana. Mol Ecol 15(6):1689–1697CrossRefPubMedGoogle Scholar
  31. 31.
    Larder BA, Kemp SD, Darby G (1987) Related functional domains in virus DNA polymerases. EMBO J 6(1):169–175PubMedPubMedCentralGoogle Scholar
  32. 32.
    Plumb JA, Wright LD, Jones VL (1973) Survival of channel catfish virus in chilled, frozen, and decomposing channel catfish. Progress Fish Culturist 35:170–172CrossRefGoogle Scholar

Copyright information

© Springer-Verlag Wien 2016

Authors and Affiliations

  • Chimoné S. Dalton
    • 1
  • Karen van de Rakt
    • 1
  • Åsa Fahlman
    • 2
  • Kathreen Ruckstuhl
    • 3
  • Peter Neuhaus
    • 3
  • Richard Popko
    • 4
  • Susan Kutz
    • 1
    • 5
  • Frank van der Meer
    • 1
  1. 1.Department of Ecosystem and Public Health, Faculty of Veterinary MedicineUniversity of CalgaryCalgaryCanada
  2. 2.Department of Clinical Sciences, Faculty of Veterinary MedicineSwedish University of Agricultural SciencesUppsalaSweden
  3. 3.Department of Biological Sciences, Faculty of ScienceUniversity of CalgaryCalgaryCanada
  4. 4.Environment and natural Resources Sahtú RegionNorman WellsCanada
  5. 5.Canadian Wildlife Health Cooperative, Alberta Node, Faculty of Veterinary MedicineUniversity of CalgaryCalgaryCanada

Personalised recommendations