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In this study, we showed that the expression of cry and cp4-epsps genes in wild cotton altered the secretion of EFN, the associations with different ant species, and the levels of herbivore damage on target plants. Wcry constantly maintained a high production of EFN, regardless of the MeJA treatment, but nectar production was minimal in Wcp4-epsps. These changes in nectar inducibility seem to modify the composition of ant communities, foster the dominance of the generalist and defensive species C. planatus in Bt plants and the presence of ants without defensive role, M. ebeninum, in the herbicide tolerant genotype, while W plants had both defending species (C. planatus, C. rectangularis aulicus and P. gracilis) and invasive ant species (P. longicornis) in the same proportion. Furthermore, herbivore damage and its associated ant community were different according to the introgressed transgene.
Wild and introgressed cotton do not display phenotypic equivalence in natural conditions
In general, it has been assumed that introgressed and wild genotypes should display similar phenotypes in the absence of the selection agents targeted by transgenes. However, when we compared the control group and the three genotypes, we registered different nectar secretion patterns among them (Fig. 1). Similar results have been registered in populations of bt rice and glyphosate-tolerant sunflowers living in natural conditions where introgressed plants are different from their wild relatives5.
Transgene expression modified indirect induced defences in wild cotton
Most plants are able to induce responses after herbivore damage and/or phytohormone exogenous application (i.e. jasmonic acid, JA; methyl jasmonate, MeJA; and salicylic acid, SA)11,28,29. However, unlike wild plants without transgenes, individuals with transgenes were not sensitive to the induction treatment with MeJA for increasing their EFN production (Fig. 1). These results contrast with previous reports on cultivated varieties, such as Bt and glyphosate-resistant (cp4-epsps), in which direct defences such as gossypol terpenoids (160%), hemigossypolone (160%), helicoids 1|4 (213%) and indirect defenses, such as volatile compounds (VOCs) (171.2%) and extrafloral nectar (EFN) (133%), were reported to increase in plants sprinkled with JA and MeJA21,28,29,30.
The inability of plants with transgenes to have the production of extrafloral nectar induced in them was related to different processes dependent on the identity of the transgenes in question. Whereas Wcry control plants had a high EFN production equivalent to the induced state of W plants, EFN production in Wcp4-epsps plants was inhibited. Contrasting these findings with results obtained under controlled conditions (i.e. greenhouse and crop conditions)3,21, we suggest that EFN production is linked to genotypes with transgenes and abiotic stress in the coastal dunes, because transgenes are connected to main metabolic pathways that respond to stressful conditions21.
Wild cotton with cp4-epsps
In the absence of herbicides acting as a selection agent, wild plants with cp4-epsps exhibited large differences compared to wild plants without them. Their low nectar production ( > 8 µg/mL) (Fig. 1) could be linked to the crosstalk between the jasmonate and the salicylate (SA) pathways (Fig. 4, orange and purple section). In G. hirsutum and other species, SA signalling has been proven to negatively affect JA signalling (e.g. Zea mays, Solanum lycopersicum, Nicotiana tabacum and Arabidopsis thaliana)31,32,33: therefore, we suggest an interference between the SA and JA pathways given previous reports that an over-expression of the cp4-epsps gene modifies the second part of the shikimate pathway (post-chorismate), which leads to the synthesis of essential amino acids as phenylalanine, tryptophan, or tyrosine, the latter being a precursor of benzoic acid BE, and SA34,35 (Fig. 4, purple section). This evidence highlights that hidden crosstalk effects among different metabolic pathways can scale up and modify plant phenotypes (e.g. extrafloral nectar production).
Figure 4

A diagram illustrating how the expression of cry (A) and cp4-epsps (B) in absence of their selection agent (pests and glyphosate) can affect the extrafloral nectar production. The extrafloral nectar (EFN) production is an induced defence that can be triggered by foliar herbivory, mechanical damage, and exogenous application of phytohormones (i.e. jasmonic acid, methyl jasmonate, and salicylic acid). These factors activate the octadecanoid pathway, and therefore, the production of extrafloral nectar, (A) aqua rectangle. The (C) section is an example of this reaction in a wild cotton plant (without transgenes). After damage, the key genes (yellow mesh) of the octadecanoid pathway are activated and produce extrafloral nectar. Another scenario is when the wild cotton expresses cry genes (A section), in this case, the key genes of the octadecanoid pathway interact synergistically with the cry transgene (green mesh). This triggers an over-expression of the production of EFN (aqua thick arrow), switching from inducible to constitutive responses. When the plants express cp4-epsps (B section), the production of extrafloral nectar is reduced or inhibited. A possible answer is an over-expression of the epsps gene (gold curve arrow), that increased production of salicylic acid which creates a crosstalk between shikimate and octadecanoid pathways (black cross-talk arrow). When the shikimate pathway is activated, the principal inducible defence is the production of volatile organic compounds (VOCs) (pink rectangle).

Full size image

Wild cotton with cry
Wild cotton plants with cry genes continuously produced EFN as a constitutive defence (Fig. 1), in equivalent quantities as the induced state of W plants. EFN production is regulated by the octadecanoid signalling pathway, which can be activated by herbivore damage, mechanical damage, and phytohormones, such as JA and MeJA21,28 (Fig. 4, green section). However, for cotton, a specific elicitor is not necessary36. Four key genes for the synthesis of JA and MeJA have been described: AOS, AOC, HPL, and COI137. In Bt maize, studies comparing GM corn and its isogenic lines report an increase of 24% in phenols and 63% of DIMBOA (2,4-dihidroxi-7-metoxi-1,4-benzoxazin-3-ona; natural defences against lepidopteran herbivores)11. This is consistent with observations of a synergy between maize direct defences and Bt genes, after exogenous applications of JA (Fig. 4, orange section). Considering the latter, we suggest that Wcry cotton may present a similar response, as the genes activating the JA pathway are GhAOS and GhCOI1 (homologs to maize JA biosynthesis genes: ZmAOS and ZmCOI1), in addition to Ghppo1, which confers natural resistance to lepidopteran pest, such as H. armigera38. The interaction of cry with other genes could modify the production of EFN in Wcry plants.
Effect of the transgenes’ expression on ants associated to wild cotton
We identified eight species of ants harvesting EFN (Table 2), but with distinctive communities as a function of the plant genotype. This result suggests that the change in quantity, and possibly the composition and quality of EFN, can influence the ant community associated with G. hirsutum39,40,41.
Changes in plant reward production could potentially compromise the attraction of natural enemies of herbivores42. In our study, the availability of EFN was modified. Although species richness was the same as in W plants (Table 2), the most abundant ant species associated with Wcp4-epsps plants, M. ebeninum, is considered a generalist species. Moreover, due to the lack of aggressive behaviour, this species does not represent an effective biotic defence43. The high abundance of this non-defensive species could be associated with the greater herbivore damage observed in Wcp4-epsps plants (Fig. 2). In contrast, W or Wcry plants showed a greater abundance of more aggressive ant species such as C. planatus, C. rectangulatus, and P. brunneus and significantly less herbivore damage.
In Wcry cotton, the community of patrolling ants was mainly dominated by C. planatus, in both treatments (control and induction). Interestingly, although the amount of nectar did not vary between treatments, the abundance of ants was significantly different. The dominance of a single ant species could have benefited the plants with increased indirect defence, reducing herbivore damage and promoting a greater seed production per plant, as described in Turnera ulmifolia44, Schomburgkia tibicinis45, and Opuntia stricta42. However, considering the aggressive and dominant behaviour of C. planatus, there may be ecological costs through antagonistic relationships with pollinators. Ants can interrupt pollination and affect plant fitness25,46,47. The outcome of these mutualistic and antagonistic interactions requires further study.
Effects of transgenes on herbivore damage
Considering that the type of mutualism that cotton sustains with ants is defensive, we suggest that the change we observed in the composition of ants is likely to have influenced herbivore damage in the different genotypes, which in turn has the potential to reduce fitness as shown by other studies of cotton48,49,50. However, a study carried out on wild upland cotton reported that plants tolerate intermediate levels of leaf damage inflicted by leaf-chewing insects ( More

Sample collection
Algae were sampled from August 27th to September 21st (2017) from seven populations also collected for Bonthond et al. [28], including three native populations; Akkeshi (Japan), Soukanzan (Japan), Rongcheng (China); and four non-native populations; Pleudihen-sur-Rance (France), Nordstrand (Germany), Cape Charles Beach (Viriginia) and Tomales Bay (California, Fig. 1, Table S1). Individuals fixed to hard substratum (see [30]) were sampled at least a meter apart from one another and stored in separate plastic bags. As A. vermiculophyllum has a complex, haplodiplontic life-cycle only diploids were included in the experiment. Life-cycle stages were identified in the field with a dissecting microscope or post-hoc by microsatellite genotyping [31]. After transport in coolers and storage at 4 °C in the lab, bags with algae were shipped to Germany, arriving within 4–6 days after collection. In the climate room (15 °C), individuals were transferred to separate transparent aquaria with transparent lids, containing 1.75 L artificial seawater (ASW) prepared from tap water and 24 gL−1 artificial sea salt without CaCO3 (high CaCO3 concentrations increase disease risk, Weinberger data unpublished) and exposed to 12 h of light per day (86.0 µmol m−2s−1 at the water surface). Aquaria were moderately aerated with aeration stones. Per population, four diploid individuals were acclimated over 31–32 days to climate room conditions prior to starting the experiment. Water was exchanged weekly with new ASW enriched with 2 mL Provasoli-Enrichment Solution (PES; [32]). At the start of the experiment, wet weight was recorded and individuals were divided into two parts of ~10 g each and placed into two plastic tanks with 1.75 L water and 2 mL PES (Fig. 1).
Fig. 1: Schematic overview of the sampling design and experimental process.

Algae were collected from native populations Rongcheng (ron), Soukanzan (sou) and Akkeshi (akk) and non-native populations Tomales Bay (tmb), Cape Charles Beach (ccb), Pleudihen-sur-Rance (fdm) and Nordstrand (nor). In the climate room algae were acclimated for 5 weeks and divided into two thalli. One of the thalli was treated for three days with an antibiotic mixture after which both groups were monitored for six weeks, during which the treated algae received inoculum with each water change. Microbiota samples were taken in the field (tfield), directly after disturbance (t0) and after 1, 2, 4 and 6 weeks (t1, t2, t4 and t6).

Full size image

Experimental setup
To rigorously disturb the microbial community, one of each of the pairs of aquaria containing the same algal individual was treated with a combination of antibiotics, aiming to increase the effectivity (10 mgL−1 ampicillin, 10 mgL−1 streptomycin, 10 mgL−1 chloramphenicol) and the other (control) remained untreated. All experimental work was conducted with disposable gloves and sterilized equipment, to minimize contamination. After three days, the water was removed from all tanks (treated and control) and the wet weight was recorded for all algae. All individuals were rinsed with one 1.75 L volume ASW and re-incubated in 1.75 L ASW. Subsequently, both groups received new ASW with 2 mL PES weekly and individuals treated with antibiotics received also 2 mL inoculum. The inoculum was prepared from individuals of all 7 populations, following the procedure to remove epibiota as described in Bonthond et al. [28]. Briefly, apical fragments of 1 g were separated from the thallus and transferred to 50 mL tubes containing 15 ± 1 glass beads (3 mm) and 15 mL ASW and vortexed for 6 min to separate epibiota from the algal tissue. In total, 8 samples were prepared from one individual per population. The resulting suspensions were pooled and mixed with glycerol (20% final glycerol concentration), aliquoted in 50 mL tubes and stored at −20 °C. For each water exchange, a new aliquot was defrosted at room temperature and added to the water of treated algae. Wet weight was recorded weekly with water exchanges. Before weighing the individual on aluminum foil, it was dipped twice on a separate aluminum foil sheet, to reduce attached water in a systematic way. Endo- and epiphytic microbiota were sampled in the field (tfield, [28]), at the start of the experiment (t0), after one week (t1), two weeks (t2), four weeks (t4) and six weeks (t6, Fig. 1). To equalize acclimation times across populations the experiment was stacked into five groups (Table S2). At each sampling moment, 0.5 or 1 g of tissue was separated from all individuals with sterilized forceps and epibiota were extracted similarly to the preparation of the inoculum. The resulting suspension was filtered through 0.2 µm pore size PCTA filters. Both the filters and the remaining tissue were preserved at −20 °C.
DNA extraction and amplicon sequencing
Tissue samples were defrosted, rinsed with absolute ethanol and DNA free water to remove hydro- and moderately lipophilic cells and molecules from the surface and cut to fragments with sterilized scissors. DNA was then extracted from these fragments (endobiota) and from preserved filters (epibiota) using the ZYMO Fecal/soil microbe kit (D6102; ZYMO-Research, Irvine, CA, USA), following the manufacturer’s protocol. Although this method to separate endo- and epibiota was shown to resolve distinct communities [28], tightly attached epiphytic cells may not be completely removed from the surface and detectable in endophytic samples as well. Two 16S-V4 amplicon libraries, over which the samples were divided in a balanced manner, were prepared as in Bonthond et al. [28], following the two-step PCR strategy from Gohl et al. [33], using the same set of 16S-V4 target primers and indexing primers. The libraries were sequenced on the Illumina MiSeq platform (2×300 PE) at the Max-Planck-Institute for Evolutionary Biology (Plön, Germany), including four negative DNA extraction controls and four negative and positive PCR controls (mock communities; ZYMO-D6311). The fastq files were de-multiplexed (0 mismatches). Relevant field samples from Bonthond et al. [28] were combined with the new dataset and assembled, quality filtered and classified altogether with Mothur v1.43.0 [34] using the SILVA-alignment release 132 [35]. Sequences were clustered within 3% dissimilarity into OTUs using the opticlust algorithm. Mitochondrial, chloroplast, eukaryotic and unclassified sequences were removed. To prepare the community matrix we discarded singleton OTUs (in the full dataset), samples with More

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Department of Geography, Texas A&M University, College Station, TX, USA
J. Loisel

Department of Geography, University of Exeter, Exeter, UK
A. V. Gallego-Sala, M. J. Amesbury, D. J. Charman & T. P. Roland

Ecosystems and Environment Research Programme, University of Helsinki, Helsinki, Finland
M. J. Amesbury, A. Korhola, M. Väliranta, S. Juutinen, K. Minkkinen & S. Piilo

Department of Geography and Geotop Research Center, University of Quebec at Montreal, Montreal, Quebec, Canada
G. Magnan & M. Garneau

Magister of Environment and Soil Science Department, Tanjungpura University, Pontianak, Indonesia
G. Anshari

Department of Geography and Environment, University of Hawaii at Manoa, Honolulu, HI, USA
D. W. Beilman

Department of Ecology and Territory, Pontificial Xavierian University, Bogota, Colombia
J. C. Benavides

Organic Geochemistry Unit, School of Chemistry, and School of Earth Sciences, University of Bristol, Bristol, UK
J. Blewett & B. D. A. Naafs

Environmental Studies Program and Earth and Oceanographic Science Department, Bowdoin College, Brunswick, ME, USA
P. Camill

Department of Geology, Chulalongkorn University, Bangkok, Thailand
S. Chawchai

Department of Geography, University of California, Los Angeles, Los Angeles, CA, USA
A. Hedgpeth

Max Planck Institute for Meteorology, Hamburg, Germany
T. Kleinen & V. Brovkin

Faculty of Engineering, Chemical and Environmental Engineering, University of Nottingham, Nottingham, UK
D. Large

Centro de Investigación GAIA Antártica, University of Magallanes, Punta Arenas, Chile
C. A. Mansilla

Climate and Environmental Physics, Physics Institute and Oeschger Centre for Climate Change Research, University of Bern, Bern, Switzerland
J. Müller & F. Joos

Consortium Érudit, Université de Montréal, Montreal, Quebec, Canada
S. van Bellen

Department of Ecology and Conservation Biology, Texas A&M University, College Station, TX, USA
J. B. West

Department of Earth and Environmental Sciences, Lehigh University, Bethlehem, PA, USA
Z. Yu

Institute for Peat and Mire Research, School of Geographical Sciences, Northeast Normal University, Changchun, China
Z. Yu

Department of Environmental Studies, Mount Holyoke College, South Hadley, MA, USA
J. L. Bubier

Department of Geography, McGill University, Montreal, Quebec, Canada
T. Moore

Department of Physical Geography, Stockholm University, Stockholm, Sweden
A. B. K. Sannel

School of Geography, Geology and the Environment, University of Leicester, Leicester, UK
S. Page

Department of Earth and Environmental Sciences, KU Leuven, Leuven, Belgium
M. Bechtold & W. Swinnen

School of Geography & Sustainable Development, University of St Andrews, St Andrews, UK
L. E. S. Cole

Department of Earth, Ocean & Atmospheric Science, Florida State University, Tallahassee, FL, USA
J. P. Chanton

Department of Bioscience, Aarhus University, Roskilde, Denmark
T. R. Christensen

Department of Earth Sciences, University of Toronto, Toronto, Ontario, Canada
M. A. Davies & S. A. Finkelstein

Instituto Franco-Argentino para el Estudio del Clima y sus Impactos, Buenos Aires, Argentina
F. De Vleeschouwer

Institute for the Study of Earth, Oceans, and Space, University of New Hampshire, Durham, NH, USA
S. Frolking & C. Treat

Department of Geobotany and Plant Ecology, University of Lodz, Lodz, Poland
M. Gałka

Laboratoire d’Ecologie Fonctionnelle et Environnement, UMR 5245, CNRS-UPS-INPT, Toulouse, France
L. Gandois

Cranfield Soil and Agrifood Institute, Cranfield University, Cranfield, UK
N. Girkin

Department of Renewable Resources, University of Alberta, Edmonton, Alberta, Canada
L. I. Harris

Stockholm Environment Institute, University of York, York, UK
A. Heinemeyer

Max Planck Institute for Biogeochemistry, Jena, Germany
A. M. Hoyt

Lawrence Berkeley National Laboratory, Berkeley, CA, USA
A. M. Hoyt

Florence Bascom Geoscience Center, United States Geological Survey, Reston, VA, USA
M. C. Jones

Department of Marine and Coastal Environmental Science, Texas A&M University at Galveston, Galveston, TX, USA
K. Kaiser

Department of Biology, University of Victoria, Victoria, British Columbia, Canada
T. Lacourse

Faculty of Geographical and Geological Sciences, Climate Change Ecology Research Unit, Adam Mickiewicz University, Poznań, Poland
M. Lamentowicz

Natural Resources Institute Finland (Luke), Helsinki, Finland
T. Larmola

Agroscope, Zurich, Switzerland
J. Leifeld

Institute for Atmospheric and Earth System Research, University of Helsinki, Helsinki, Finland
A. Lohila

Finnish Meteorological Institute, Climate System Research, Helsinki, Finland
A. Lohila

Department of Geography, Royal Holloway, University of London, Egham, UK
A. M. Milner

Department of Forest Sciences, University of Helsinki, Helsinki, Finland
K. Minkkinen

School of Earth and Environmental Sciences, University of Queensland, Brisbane, Queensland, Australia
P. Moss

Lamont-Doherty Earth Observatory, Palisades, NY, USA
J. Nichols

National Park Service, Washington DC, WA, USA
J. O’Donnell

Department of Environment & Geography, University of York, York, UK
R. Payne

Department of Chemistry, and Department of Geological and Environmental Science, Hope College, Holland, MI, USA
M. Philben

Department of Geography and Environmental Science, University of Reading, Reading, UK
A. Quillet

Department of Applied Earth Sciences, Uva Wellassa University, Badulla, Sri Lanka
A. S. Ratnayake

School of Biosciences, University of Nottingham, Nottingham, UK
S. Sjögersten

Département de Géographie, Université de Montréal, Montréal, Québec, Canada
O. Sonnentag & J. Talbot

Geography, School of Natural and Built Environment, Queen’s University Belfast, Belfast, UK
G. T. Swindles

Department of Environmental Science, Policy, and Management, University of California, Berkeley, Berkeley, CA, USA
A. C. Valach

Department of Environment and Sustainability, Grenfell Campus, Memorial University, Corner Brook, Newfoundland, Canada
J. Wu More

Collection of wild carp from a CyHV-3-exposed population
This study was carried out in accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. All protocols for sampling, procedures and experimental endpoints involving live fish conducted in this study were approved by the Institutional Animal Care & Use Committee (IACUC), University of Minnesota (St. Paul, Minnesota, USA), under the approval numbers IACUC-1806-36036A and 1808-36276A. Experiments were performed in compliance with the ARRIVE guidelines on animal research32.
Wild carp were sampled from Lake Elysian (Waseca County, Minnesota, Coordinates: 44.178144, − 93.69066) by boat electrofishing from September 3 to 9, 2019 (Fig. 1a). This lake was expected to have a CyHV-3-exposed carp population following a confirmed outbreak in 20173. Captured wild adult carp (n = 116) were euthanized by immersion in a solution of ~ 3 mL/L pure clove oil (90% Eugenol; Velona, Elk Grove Village, IL, USA) for 15 min and transported on ice to the University of Minnesota for necropsy. Brain, gill and kidney tissues from up to three carp were pooled in a 1:5 (weight:volume) dilution of Hank’s Balanced Salt Solution (HBSS; Cellgro, Lincoln, NE, USA) containing 100 IU/mL of Penicillin and Streptomycin and maintained at a pH of 7.4 at 4 °C for 24 h prior to preparation for qPCR and cell culture screening for CyHV-3 (described below). Gill tissues from ten freshly-dead carp obtained from a shallow bay in the Southern portion of the lake were also obtained and pooled by five individuals for a total of two sample pools.
Figure 1

(a) Generated using ArcMap (v10.8.1, https://desktop.arcgis.com/en/arcmap/), shows the approximate locations of sampling effort and mortality observations on Lake Elysian. Bathymetric contours indicate depth in 5 ft increments. (b, c) Pathology of a representative individual wild carp sampled from Lake Elysian. Arrows on (b, c) denote frayed fins (vermillion), loss of mucosal layer (white), loss of scales and epidermis (black), enopthalmia (bluish green), gill necrosis (sky blue).

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An additional 17 wild carp collected as part of the previously described sampling event were placed in an aerated live well and transferred to the Minnesota Aquatic Invasive Species Research Center’s Containment Laboratory (MCL). These carp were housed in a ~ 1400 L tank with flow through well water at 20 °C and treated with 0.6% aquarium salt once per day. Carp were acclimated for 1 day and then anesthetized via immersion in a solution of 100 µL/L of clove oil and uniquely marked using colored injectable elastomer (Northwest Marine Technology, Anacortes, WA, USA). Additionally, a small portion (~ 0.2 cm2) of each carp’s gills were sampled for qPCR screening for CyHV-3 and tested immediately. Carp determined to be CyHV-3 negative (n = 12) were euthanized following testing. Carp determined to be CyHV-3-positive (n = 5) by specific qPCR were held for a total of 5 days, during which, water temperature was gradually increased to 26 °C in order to increase viral shedding. CyHV-3-positive carp gill biopsies were again sampled and screened on the fifth day to identify carp with high qPCR copy numbers. All CyHV-3-positive carp were then euthanized, and the brain, gill and kidney tissues were removed as previously described. Pooled tissues from two wild carp with clinical signs consistent with KHVD (Fig. 1b,c) and with high qPCR copy numbers, were subjected to cell culture immediately following necropsy. In addition, a 10 g portion of this pooled tissue was processed and used to challenge naive carp in the in-vivo infection model. Tissues were homogenized in a 1:5 volume of HBSS containing 100 IU/mL Penicillin and Streptomycin (pH = 7.4). The sample was centrifuged at 2360 × g at 25 °C for 10 min, then the supernatant was passed through a 0.45 µm syringe filter.
In-vivo infection trial
To increase the potential of obtaining an isolate of CyHV-3, naïve carp previously determined to be CyHV-3 negative by qPCR, were challenged with CyHV-3-positive tissue homogenates obtained from wild carp. Two naïve carp, purchased from Osage Catfisheries (Osage Beach, MO, USA), were pair housed in a 60 L aquarium with flow through well water (flow rate = 3–4 tank volumes/h) at 21–22 °C. Aquaria were set up with a standpipe drain covered by a cylindrical wire screen filter of approximately 15 cm in length and 4.4 cm in diameter. Additionally, a PVC pipe section of 15 cm in length and 10 cm in diameter was added to each tank for shelter. Each carp was exposed to 0.5 mL of CyHV-3-positive tissue homogenate by IP-injection and monitored for signs of disease for 6 days and then euthanized. Pooled samples of brain, gill and kidney tissue were subjected to qPCR and cell culture analysis. Following cell culture analysis (below) a second infection trial was performed to satisfy River’s postulates (i.e. that CyHV-3 isolated from wild diseased carp would cause similar disease in naïve carp)33. Two additional naïve carp purchased from Osage Catfisheries were IP-injected with 0.5 mL of CyHV-3-positive (qPCR and cell culture positive) cell culture supernatant. Carp were housed and observed for disease signs as previously described for 11 days and then sacrificed. Pooled samples of brain, gill, and kidney then were tested by CyHV-3-specific qPCR to confirm the presence of CyHV-3.
Cell culture analysis
CCB cells were maintained in Eagle’s Minimum Essential Medium (EMEM) containing Eagles’s salts (Sigma, St. Louis, MO, USA), 10% fetal bovine serum (FBS), 1% non-essential amino acids (NEAA, Sigma), 2 mM l-glutamine and glucose (Sigma) up to 4.5 g/L. The KF-1 cells were cultured in EMEM containing Eagles’s salts (Sigma), 10% FBS and 0.025 M HEPES. Penicillin 100 U/L and streptomycin 0.1 mg/L (Sigma) were used as an anti-bacterial agent in both cell culture media and the cells were maintained at 25 °C.
Cell culture methods to isolate CyHV-3 were performed according to the US Fish and Wildlife Service and American Fisheries Society-Fish Health Section Blue Book34. Briefly, pooled tissues were homogenized in Hank’s Balanced Salt Solution (HBSS; Cellgro) and centrifuged at 2360 × g for 15 min. The inoculum was added to the 24-well plates with 80% confluent cell cultures in two dilutions, (1/10 and 1/100) and incubated at 25 °C for 14 days. A blind passage was performed for an additional 14 days if no cytopathic effects (CPE) were observed on the first passage. If CPE was observed during the first passage, then the second passage was performed in a 25 cm2 flask. The virus was harvested when CPE reached 70–80% of the monolayer. The infected cultures were exposed to two freeze/thaw cycles at − 80 °C, and then centrifuged at 3800 × g for 15 min at 4 °C. The clarified supernatants and pellets were collected and stored at − 80 °C.
Whole-genome sequencing and sequence analysis
Whole-genome sequencing was performed at the University of Minnesota Veterinary Diagnostic Laboratory for genetic characterization of the CyHV-3 isolate (KHV/Elysian/2019) obtained from wild carp. In brief, after CCB cells, infected with wild carp tissues, reached 80% CPE, the supernatant was collected and stored at − 80 °C. The frozen supernatant was freeze-thawed three times, and centrifuged at 2896 × g for 25 min at 4 °C. Nucleic acid purification of CCB cell culture supernatant was done using a QIAamp MinElute Virus Spin Kit (Qiagen, Hilden, Germany) following manufacturer instructions. The extracted nucleic acids were subjected to library preparation using Nextera Flex DNA library kit (Illumina, San Diego, CA, USA) following manufacturer instructions. The library was normalized according to the median fragment size measured by Tape Station 2.0 (Agilent, Santa Clara, CA, USA) and library concentration measured by Qubit. The library was submitted to the University of Minnesota Genomic Center (UMGC) for sequencing via MiSeq V3 (2X75-bp) paired end chemistry.
Raw fastq files were trimmed to remove Illumina adapters using Trimmomatic (v 0.39) with a minimum quality score of 20. Then, bowtie2 (v 2.3.5) was used to remove host contamination and unmapped reads were used for assembly with SPAdes (v3.13.0) with k-mer values of 29, 33 and 55 with the options “careful with a minimum coverage of 5 reads per contig”. Then contigs were searched into the RefSeq viral and non-redundant protein reference databases using Diamond BLASTx with an e-value of 1e − 5 for significant hits. Taxon assignments were made with MEGAN6 Community Edition according to the lowest-common-ancestor algorithm. ORFs prediction and genome annotation were done using Prokka (v1.14.5). The resulting alignment (GenBank accession no. MT914509) was aligned with 19 other CyHV-3 genomes available on NCBI using Mafft (v7) with the FFT-NS-2 alignment strategy and the following parameters: scoring matrix BLOUSUM62, gap open penalty 1.53, offset value 0. Model selection, maximum likelihood (ML) tree construction, and calculation of bootstrap values were done with R 4.0 (R Software) using phangorn (v2.5.5). ML trees were constructed using the top scoring model (GTR + G + I) and 100 bootstrap replicates were generated to assess the reliability of clades obtained in the tree. Additionally, this genome assembly was compared with the previously reported thymidine kinase gene sequence obtained from carp sampled during a large mortality event in Lake Elysian in 2017 (F36, GenBank accession no. MK987089).
Investigation of species specificity
To investigate the host range of KHV/Elysian/2019, six carp purchased from Osage Catfisheries, previously determined to be CyHV-3-negative by qPCR, were intraperitoneally (IP) injected with 0.5 mL of the filtered tissue homogenate material (Fig. 2a). The IP-injected carp (IP-carp) were housed as previously described for 9 days prior to their use in the cohabitation trial (Fig. 2b). The IP-carp were monitored twice daily for signs of disease. After 9 days the gills, skin and vent of each IP-carp was swabbed aseptically with a single sterile cotton swab (Dynarex, Orangeburg, NY, USA) for determination of viral load by qPCR. FHM and goldfish were challenged with CyHV-3 via cohabitation. One cohabitation tank (tank A) contained ten naïve FHM, five naïve sentinel carp (S-carp) and three IP-carp (Fig. 2a). One cohabitation tank (tank B) contained ten naïve goldfish, five naive S-carp and three of the IP-carp. S-carp were included in each tank setup to act as a positive control for within-tank transmission of CyHV-3. Two additional negative control tanks with the same stocking density and conditions contained ten naïve FHM (tank D) and ten naïve goldfish (tank E), as well as eight naïve carp (confirmed to be CyHV-3-negative by specific qPCR). Average standard length and weight for fishes used in these experiments was 13 cm and 64 g for carp, 7 cm and 13 g for FHM, and 10 cm and 38 g for goldfish. All tanks consisted of ~ 60 L aquaria with flow-through well water as previously described. Fishes were fed a commercial feed (Skretting classic trout, Skretting, Tooele, UT, USA) daily and monitored twice daily to observe changes to fish health. IP-carp that died during the trial were allowed to remain in the tank for 24 h prior to removal for necropsy, but any morbidity or mortality of other experimental groups were immediately removed and necropsied.
Figure 2

(a) Shows a schematic of the cohabitation disease trial. Vermillion arrows denote inoculation of IP-carp with CyHV-3 positive tissue homogenate, blue arrows denote introduction of IP carp for cohabitation with fishes in experimental tanks, and the reddish purple arrow indicates the tissue origin of CyHV-3-positive S-carp. (b) Shows a schematic of experimental flow through chambers with black arrows indicating the direction of water flow. (c) Shows a time-line of various samples.

Full size image

At 0, 3, 6, 9, 12, and 15 days post exposure (dpe) by cohabitation, five FHM, five goldfish, and all IP-carp and S-carp from each tank were anesthetized by immersion in a buffered solution of 100 mg/L of MS-222 and the gills, skin and vent of each fish was swabbed with a sterile swab for determination of viral load by qPCR (Fig. 2c). For FHM and goldfish, the five individuals were randomly sampled at each time-point. Additionally, the wire screen filter of the outflow standpipe was swabbed at the same intervals during the course of the trial to evaluate loading of CyHV-3 DNA in the environment. All swabs were stored at − 20 °C in individual plastic bags until nucleic acid extraction could be performed. At 11 dpe, half of the FHM and goldfish from cohabitation tanks were euthanized by immersion in a buffered solution of 3 g/L of MS-222 and necropsied (Fig. 2c). The remaining FHM and goldfish were maintained until 20 dpe and then euthanized and necropsied. To visually record the presence of gross pathology, representative IP carp, and fish from cohabitation groups (S-carp, FHM, and goldfish) were randomly selected and photographed at 0 and 6 dpe in a small glass aquarium (Fig. 3).
Figure 3

Representative fishes photographed before and after exposure to CyHV-3. Note, fishes photographed at 0 dpe may not be the same individual as those at 6 dpe. dpe days post exposure via cohabitation, IP-carp intraperitoneally injected carp, S-carp cohabitated sentinel carp, FHM fathead minnow. Arrows denote frayed fins (vermillion), loss of mucosal layer (white), scale pocket edema (black). Additionally, normal morphological features of mature male fathead minnows are indicated for nuptial tubercles (bluish green), and nape pads varying in prominence (reddish purple).

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For each necropsied fish, wet mounts of gill and skin scrapes were viewed at 40× magnification to identify potential parasitic infections. Then the skin of each necropsied fish was rinsed briefly with 70% ETOH and clean water. Brain, gill, kidney and skin tissue were collected individually for each fish and split into two duplicate samples. The first sample duplicates were placed in Whirl–Pak sample bags (Nasco, Fort Atkinson, WI, USA) and preserved at − 20 °C until nucleic acid extraction and screening for CyHV-3 DNA was performed. The second sample duplicates were placed in 1 mL of RNAlater solution (Ambion) in 1.5 mL microcentrifuge tubes (Globe Scientific, Mahwah, NJ, USA) and frozen at − 20 °C. An individual FHM and goldfish from each time-point (11- and 20-dpe) was preserved in 10% NBF (TissuePro, Gainesville, FL, USA) for histological analysis. Individual representatives of each species from control tanks and moribund S-carps from each experimental tank were also preserved for histological analysis.
Due to the detection of CyHV-3 DNA in a single FHM in tank A, a second trial with FHM (tank C) was performed as described previously (Fig. 2a). Brain, gill, kidney, and skin tissue from two S-carp exposed in the first trial with disease signs and positive qPCR test for CyHV-3 (tank A) were pooled, homogenized and filtered as previously described. Three new carp purchased from Osage Catfisheries were IP injected with 0.5 mL of this tissue homogenate and maintained as previously described for 9 days prior to screening for CyHV-3 by qPCR and used in the cohabitation trial. All other conditions and procedures were done as described for the first cohabitation trial with the following exceptions. In the second trial, portions of brain, gill, kidney and skin tissues obtained from a moribund S-carp at 5 dpe and four FHM at 11 dpe, respectively, were pooled as previously described and subjected to cell culture. Additionally, duplicate swabs from the tank C outflow standpipe filter were obtained and preserved in 1 mL of RNAlater solution (Sigma) as previously described for tissue samples.
Nucleic acid purification using chelex resin and detection of CyHV-3 by qPCR
For nucleic acid purification, chelex resin (Sigma) was used as described by Zida et al.35 and briefly summarized here. For pooled tissue samples, approximately 100 mg of each tissue was homogenized in 1 mL of nuclease free water (NFW) and then centrifuged, with 50 μL of the resulting supernatant later used as starting material. For swabs, the cotton end was cut off and vortexed, then centrifuged and finally the cotton was removed leaving the supernatant. For each sample type, 150 μL of chilled 80% ETOH was added, then centrifuged and the supernatant removed. Samples were allowed to air dry for 10 min to remove residual ETOH. 150 μL of 20% Chelex was added to each sample and vortexed. Samples were then incubated at 90 °C for 20 min and centrifuged and immediately used for qPCR.
A Taqman probe-based qPCR was used for the detection of CyHV-3 DNA targeting the ORF89 gene36 using a StepOnePlus thermocycler with default settings (Applied Biosystems). Nucleic acid purifications from all samples were screened for CyHV-3 using a PrimeTime gene expression master mix kit (Integrated DNA Technologies, Coralville, IA, USA), with each reaction containing 400 nM of primers (KHV-86f: GAC-GCC-GGA-GAC-CTT-GTG, KHV-163r: CGG-GTT-GTT-ATT-TTT-GTC-CTT-GTT) and 250 nM of the probe (KHV-109p: [TAMRA] CTT-CCT-CTG-CTC-GGC-GAG-CAC-G-[IBRQ]. The reaction mix was subjected to an initial denaturation at 95 °C for 3 min, followed by 40 cycles of denaturation at 95 °C for five sec and annealing at 60 °C for 30 s. A threshold cycle of 38 was used as a cut off. The standard curve for quantification of CyHV-3 genomes was performed using a laboratory synthesized DNA fragment containing the ORF89 sequence as previously described by Padhi et al.3. The results for virus load are presented as the number of viral copies per mL of tissue supernatant. All samples obtained from FHM and goldfish were tested in triplicate with the exception of samples that had positive qPCR Ct values, which were re-tested up to six times.
RNA purification and reverse transcription polymerase chain reaction (RT-PCR)
Individual tissues of preserved brain, gill, kidney, and skin from one representative S-carp from each experimental tank (A, B and C) were selected as positive controls for CyHV-3 mRNA detection (total of 12 tissue samples). All preserved tissue samples from FHM or goldfish which had at least one positive qPCR test were also screened for CyHV-3 mRNA to determine if replicating virus was present (total of eight tissue samples). Additionally, preserved swabs of the outflow standpipe filter were also screened. For RNA purification, RNA was extracted from tissues using the RNeasy Mini Kit (Qiagen) according to the manufacturer instructions for animal tissues, using ~ 30 mg tissue samples preserved in RNAlater. For swabs, cotton was cut from the end of the swab and used as the starting material. CyHV-3 mRNA was detected using the RT-PCR developed by Yuasa et al.29 with the primers, (KHV RT F3: GCC-ATC-GAC-ATC-ATG-GTG-CA, KHV RT R1: AAT-GCC-GCT-GGA-AGC-CAG-GT). The RT-PCR was performed using a One-step RT-PCR kit (Qiagen) according to the manufacturer instructions. The reaction mix was subjected to a single step of reverse transcription at 50 °C for 30 min and denaturation at 95 °C for 15 min, followed by 40 cycles of: 94 °C for 30 s, 65 °C for 30 s, 72 °C for one minute and a final extension step was 72 °C for 10 min. PCR products were separated and visualized on 2% agarose gels containing 0.75 μg/mL ethidium bromide (Genesee Scientific, San Diego, CA, USA). PCR products for carp, FHM and goldfish templates (clear band at the 219 bp location) were cut from gels and purified by precipitation with a 20% PEG, 2.5 M NaCl solution. Purified RT-PCR products were subjected to Sanger sequencing at the University of Minnesota Genomics Center (UMGC). Sequences were trimmed and analyzed using 4 peaks (v1.8) and consensus sequences were generated using BioEdit (v7.2.1). Sequence identities were compared with available reference sequences by BLASTn analysis of the National Center of Biotechnology sequence database.
Histology
Histology was used to demonstrate the presence or absence of lesions in cohabitation disease trial specimens. Histological samples of gill tissue were prepared from formalin-fixed samples of representative fishes of each species from trial and control tanks. Gill samples were dissected from formalin-fixed specimens and decalcified in 10% ethylenediaminetetraacetic acid (EDTA) for 10 days. Following decalcification, samples were dehydrated in an ethanol series to 100% ethanol, infiltrated with toluene, and subsequently embedded in paraffin. Paraffin sections were cut at 6 µm thickness using a Leica Jung 820 Histocut Rotary Microtome and mounted on slides. Sections were stained with Hematoxylin and Eosin using a protocol modified from Humasson37.
Statistical analysis
R 4.0 (R Software) was used for statistical analysis and data presentation. CyHV-3 qPCR copy numbers are presented as averages of all positive tests for samples with duplicate tests and were Log transformed prior to statistical testing. Significant differences (p  More

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