Introduction
Environmental variability, combined with heterogeneous population structure, may lead to regional adaptations among populations (Antonovics Reference Antonovics1971). Quantifying differences among wild populations can be challenging yet provides valuable information on the process of regional differentiation and local adaptation. Common garden experiments are a powerful approach for studying local adaptations, and they help control for phenomena such as phenotypic plasticity, maternal effects, and, to a certain extent, genotype-by-environment interactions (de Villemereuil et al. Reference de Villemereuil, Gaggiotti, Mouterde and Till-Bottraud2016). However, these experiments depend on reliable access to study organisms from defined source populations, which can prevent the use of this approach for many nonmodel species.
Laboratory colonies offer ready access to healthy organisms, thus providing crucial opportunities to describe population differences within a controlled environment (Roe et al. Reference Roe, Demidovich and Dedes2018). Although laboratory colonies are an essential part of scientific research, they are challenging to develop and require careful planning. When starting new colonies, the experimental goals of the investigator will determine the appropriate breeding regime and long-term maintenance strategy for each new colony (Godinho et al. Reference Godinho, Cruz, Charlery de la Masselière, Teodoro-Paulo, Eira and Fragata2020). Different breeding regimes can dramatically influence the genetic structure of laboratory colonies, resulting in either highly diverse, outbred colonies or highly similar, inbred lines. Exploring questions about local adaptation using laboratory colonies requires maintaining representative genetic variation from multiple independent source populations (Kawecki and Ebert Reference Kawecki and Ebert2004). Maintaining high standing genetic diversity within these laboratory populations is also necessary to reflect the natural variation within wild populations, help mitigate the effects of genetic drift, and, to some extent, reduce any negative effects of artificial selection that can occur during colony establishment. Using a large number of founders for each population and in each successive generations is a crucial step to maintain high standing genetic variation and to avoid founder effects. In theory, maintaining colonies with several hundred breeding individuals for each generation is sufficient to avoid significant genetic drift for 100 generations (Berthier et al. Reference Berthier, Chapuis, Simpson, Ferenz, Kane and Kang2010).
Spruce budworm, Choristoneura fumiferana (Clemens) (Lepidoptera: Tortricidae), is the most severe defoliator of conifers in the boreal forest (MacLean Reference MacLean1980; Blais Reference Blais1983; Hardy et al. Reference Hardy, Mainville and Schmitt1986; Candau et al. Reference Candau, Fleming and Hopkin1998) and is maintained in a colony by Natural Resources Canada at the Insect Production and Quarantine Laboratory at the Great Lakes Forestry Centre in Sault Ste. Marie, Ontario, Canada. Since the colony’s inception, researchers have relied extensively on these laboratory colonies to improve our knowledge of spruce budworm biology. The existing colony was initiated in 1961 but has not received new genetic stock since the mid-1980s (Roe et al. Reference Roe, Demidovich and Dedes2018); it has also experienced at least one significant population bottleneck (van Frankenhuyzen et al. Reference van Frankenhuyzen, Ebling, McCron, Ladd, Gauthier and Vossbrinck2004). As such, this colony may have experienced genetic drift and adapted to artificial laboratory conditions, rendering it dissimilar to wild populations. Furthermore, this colony was initially established from populations in Ontario, which is a small region relative to spruce budworm’s overall distribution (Fig. 1).
Sampling locations of spruce budworm populations in 2017–2018 for colony initiation (orange circles). Black-shaded area shows spruce budworm distribution in Canada based on cumulative defoliation from 1966 to 2018.
We know that spruce budworm populations are not homogeneous. They show variation in genetic diversity (James et al. Reference James, Cooke, Brunet, Lumley, Sperling and Fortin2015; Blackburn et al. Reference Blackburn, Brunet, Muirhead, Cusson, Beliveau and Levesque2017; Lumley et al. Reference Lumley, Pouliot, Laroche, Boyle, Brunet and Levesque2020) and functional adaptations (Volney and Cerezke Reference Volney and Cerezke1992), whether due to environmental differences, geographic isolation (Harvey Reference Harvey1985), or postglacial colonisation history (Lumley et al. Reference Lumley, Pouliot, Laroche, Boyle, Brunet and Levesque2020). Since spruce budworm management relies on the precise quantification of survival and development rates in each population, new spruce budworm colonies derived from known locations would be a valuable resource for research on this destructive forest pest. However, establishing new colonies from wild populations is challenging. This is due, in part, to a lack of detailed methods for establishing wild spruce budworm into a laboratory colony and a loss of technical expertise. This includes the process of removing pathogens from the founding stock. Wild populations contain a variety of diseases, including pathogenic fungi, bacteria, and viruses (Ebling and Demidovich Reference Ebling and Demidovich2015). These pathogens can severely impact reared insects and can lead to high levels of mortality and subsequent colony collapse if not adequately managed (van Frankenhuyzen et al. Reference van Frankenhuyzen, Ebling, McCron, Ladd, Gauthier and Vossbrinck2004).
To address questions of local adaptation in spruce budworm, we established new laboratory colonies from defined populations. These colonies will facilitate functional comparisons within and between populations, as well as with existing strains, to understand the breadth of phenotypic variability expressed within a species. Specifically, we describe how we established new laboratory colonies from wild budworm populations. We detail how we sampled insects from host branches, established this first generation on artificial diet, eliminated pathogens from each strain, and reared subsequent generations to create new disease-free spruce budworm laboratory colonies.
Methods
Field collections
Spruce budworm is univoltine, with adults eclosing in mid-summer to lay eggs on spruce (Picea) and fir (Abies) foliage. When the eggs hatch, the first-instar larvae do not feed but immediately seek out overwintering locations and construct silken overwintering structures called hibernacula. The larvae moult within the hibernacula and then overwinter as second-instar larvae. Larvae emerge in the spring and resume development, feeding on the expanding buds of their host plants and completing their development by early summer.
Our initial field collections targeted overwintering second-instar larvae. This stage is the most transportable because second-instar larvae exist in a state of dormancy. We collected branches infested with overwintering larvae from five sites across Canada (Fig. 1; Table 1), in regions with known spruce budworm outbreaks. We relied on local collaborators to collect branches from our sampling sites. As such, sampling time depended on their availability and their ability to access the sampling sites. We targeted forest regions that had recently experienced moderate to high levels of defoliation to increase the probability of sampling enough individuals to initiate a colony. Branch tips of varying lengths were collected using pole or hand pruners, trimmed to 100-cm lengths, then placed in canvas bags. Total branch sampling varied for each location and depended on the predicted larval density. Hence, the samples consisted of 60–150 branches from each sampling location. The bags were then shipped to the Great Lakes Forestry Centre. The branches were kept cool during transport (< 5.0 °C) to reduce the risk of second-instar larval emergence in transit. Upon arrival, the branches were stored at 4 °C until processing.
Collection information for six spruce budworm populations used to establish five laboratory colonies. Larval host as follows: Sw, white spruce, Picea glauca Moench (Piceae); Sb, black spruce, Picea mariana Miller (Piceae); Fb, balsam fir (Abies balsamea Linnaeus) (Pinaceae). Colony codes assigned as per instructions in Roe et al. (Reference Roe, Demidovich and Dedes2018). NWT, Northwest Territories; AB, Alberta; ON, Ontario; NB, New Brunswick; QC, Québec; all locations are in Canada.
Flushing methods: description and comparison
Our objective was to collect postdiapause spruce budworm larvae from the branch tips as they emerged from their hibernacula (hereafter termed “flushing”). We compared two previously described techniques for flushing larvae from branches: pyramid boxes and paper cones (Miller Reference Miller1957, Reference Miller1958; Sanders Reference Sanders1980; Weber et al. Reference Weber, Volney and Spence1999).
Pyramid boxes
Pyramid boxes are free-standing enclosures containing an inner tube with entrance holes that terminated in a transparent collection jar mounted outside the top of the box (Fig. 2). Because postdiapause spruce budworm larvae are positively phototactic, the pyramid boxes are designed to collect emerging larvae that migrate towards the light within the collection jar (Miller Reference Miller1958). We cut three to five 1-m-long branches into smaller pieces, placed these inside each box, and temporarily sealed it with tape. We stored the boxes in a warm, bright room (24:0 Light:Dark; 25 ± 1 °C; 60–70% relative humidity) and checked the collecting jars daily. All collected larvae were transferred to fresh McMorran diet (McMorran Reference McMorran1965), the standard diet for spruce budworm, using a fine camel-hair paintbrush (Princeton Lauren Series 4350, Round Size 0; Princeton Artist Brush Co., Princeton, New Jersey, United States of America) to continue development.
Pyramid design for flushing overwintering spruce budworm from branches.
Paper cone method
We wrapped branches in “cones” of paper towel and collected larvae as they emerged. Specifically, we trimmed each branch to approximately 40 cm long and created bundles of three to five branches using wire coated in Tangle Foot (Scotts Miracle-Gro Company, Marysville, Ohio, United States of America; Miller Reference Miller1957; Fig. 3). Once bundled, we wrapped the branches in paper towel to form a cone that was constricted at the top and open at the bottom to encourage larvae to move downwards after emergence. Each branch bundle was hung in a separate cage above shallow pans (39 cm × 32 cm × 12 cm) containing 5 cm of water. Cages were installed within a walk-in environmental chamber (Hotpack Canada Ltd., Waterloo, Ontario, Canada) and maintained under the same conditions as the pyramid method (24:0 Light:Dark; 23 ± 1 °C; 60–70% relative humidity). We misted the bundles daily to slow desiccation and promote second-instar larval emergence. As larvae emerged from their hibernacula and “spun-down” on fine silken threads, they were collected from the paper cone or from the surface of the water. We recorded the number of larvae emerging from each bundle daily. To reduce the risk of mixing populations, we did not place branches from different populations in adjacent cages. As larvae emerged, we transferred individuals to artificial diet, as previously described. We cleaned the water pans weekly to clear away tree needles, improve larval detection, and minimise mould growth.
Individual cages, each containing a paper cone for flushing overwintering larvae from branches. Individuals cages are 45 cm × 38 cm × 61 cm (height × width × depth). Inset shows a single paper cone suspended above a collection pan.
Rearing methods: rearing protocol, single-pair mating, and quality control
We followed the standard Insect Production and Quarantine Laboratory spruce budworm-rearing protocol and quality control techniques to rear our first generation (F0) from wild-caught individuals (Ebling and Demidovich Reference Ebling and Demidovich2015). However, we included two modifications: (1) we reared larvae individually in 22-mL (0.75-oz.) cups rather than in groups, and (2) we added Fumagillin-B (4000 ppm; Medivet Animal Health, Mandeville, Louisiana, United States of America) to the artificial diet. Both modifications were adopted to manage and eliminate disease from the wild colonies, particularly microsporidia (van Frankenhuyzen et al. Reference van Frankenhuyzen, Ebling, McCron, Ladd, Gauthier and Vossbrinck2004). We reared the larvae in a walk-in environmental chamber (16:8 Light:Dark; 20 ± 1 °C; 60–70% relative humidity) (Hotpack Canada Ltd.), transferring larvae to fresh diet every 14 days until pupation. We discarded dead, diseased, and parasitised larvae. We determined the sex of pupae by counting the number of segments on the abdomen and then stored them individually until eclosion.
To produce our F1 generation, we used single-pair or small-group matings of F0 adults, rather than mass matings, in order to manage disease transmission (van Frankenhuyzen et al. Reference van Frankenhuyzen, Ebling, McCron, Ladd, Gauthier and Vossbrinck2004). As the F0 adults eclosed, we paired them with one or two members (based on availability) of the opposite sex in a 177-mL (6-oz.) cup with a paper lid (Polar Pak Company, Brampton, Ontario, Canada), provided several small strips of waxed paper as an oviposition substrate, and misted them daily with sterile water (Fig. 4). After five to seven days, we removed the wax paper strips and the attached eggs and screened the adults for pathogens. We kept only eggs that were produced by disease-free parents (see below) and discarded all other eggs. We transferred the wax strips to emergence pans, with cotton gauze (mesh size 50, Fabricland Canada, Toronto, Ontario, Canada) embedded on Parafilm (Bemis Manufacturing Company, Neenah, Wisconsin, United States of America) on the top and bottom of each pan to provide sites for hibernacula construction. Emergence pans were held for 13 days at rearing conditions (16:8 Light:Dark; 20 ± 1 °C; 60–70% relative humidity) until all viable eggs had hatched and the larvae established hibernacula. We maintained the larvae at rearing conditions for at least two weeks before placing them in diapause conditions (0:24 h Light:Dark; 2 ± 3 °C; 60–70% relative humidity) for 24–34 weeks (Ebling and Demidovich Reference Ebling and Demidovich2015).
Single-pair mating with wax paper as a substrate for laying eggs.
We screened F0 adults for pathogens, including cytoplasmic polyhedrosis virus, nuclear polyhedrosis virus, entomopoxvirus, microsporidia, fungi, and bacteria, based on methods developed by van Frankenhuyzen et al. (Reference van Frankenhuyzen, Ebling, McCron, Ladd, Gauthier and Vossbrinck2004) and Ebling and Demidovich (Reference Ebling and Demidovich2015). After we homogenised each adult in reverse osmosis water, we smeared a small sample of homogenate on a microscope slide. We used diagnostic staining with bromophenol blue and naphthalene black to identify pathogens (Evans and Shapiro Reference Evans and Shapiro1997; Fuxa et al. Reference Fuxa, Sun, Weidner and LaMotte1999; Inglis and Sikorowski Reference Inglis and Sikorowski2009). When we detected a pathogen, we quantified infection severity by recording the number of pathogen bodies in each of 20 fields of view and computed the mean number of bodies (x). We then rated infection severity as either high (x > 50), medium (5 < x < 50), low (x < 5), or negative (x = 0).
Results and discussion
Species with large geographic ranges often show regional adaptations to local environmental conditions (Chevin et al. Reference Chevin, Lande and Mace2010; see also Du et al. Reference Du, Warner, Langkilde, Robbins and Shine2010). Spruce budworm is a widely distributed forest pest and shows regional differences in both life history traits (e.g., Harvey Reference Harvey1983; Volney and Fleming Reference Volney and Fleming2007) and genomic population structure (Lumley et al. Reference Lumley, Pouliot, Laroche, Boyle, Brunet and Levesque2020). These regional differences could impact budworm management tactics and reduce their effectiveness in different parts of its range. By developing colonies of the insect from known locations, we will be able to compare populations and explore regional adaptations within this pest (Volney and Cerezke Reference Volney and Cerezke1992), a critical first step towards developing regionally informed modelling and management approaches.
Larval collections
We found that the flushing methods differed in their success and efficiency at extracting second-instar larvae from branches. We initially tested both flushing methods on a subset of branches from Ontario and Québec before selecting a method for the remaining populations. We were able to collect more larvae from Ontario branches using paper cones (32 ± 22 larvae cage−1, n = 20 cages, mean ± 1 standard deviation) than using pyramid boxes (14 ± 7 larvae cage−1, n = 39 cages), with a total of 657 larvae from cones compared to 270 larvae from boxes. We observed the same pattern for branches collected from Québec (pyramid boxes: 1 ± 1 larvae cage−1, n = 70; paper cones: 5 ± 14 larvae cage−1, n = 104), with very few larvae from the boxes compared to larvae from the cones (15 versus 730 larvae). Overall, the paper cone method was the most efficient and straightforward method to obtain enough insects to initiate a laboratory colony (Table 2). The paper cones allowed us to use slightly larger branch bundles than the enclosed pyramid boxes did, which improved flushing efficiency. This partially explains the higher larval counts that we observed. We found that this method was easier to set up, inspect for larvae, and control for humidity and airflow. However, the method required a great deal of space (Fig. 2) and the construction of individual rearing cages in a dedicated environmental chamber. The collecting trays also required frequent cleaning, and larval collecting from the cones was more time-consuming than from the pyramid boxes.
Summary of second-instar spruce budworm emergence from six wild populations flushed in paper cones. See text for description of paper cone method. NWT, Northwest Territories; AB, Alberta; ON, Ontario; NB, New Brunswick; QC, Québec; all locations are in Canada.
Initially, we expected to favour the pyramid boxes over the paper cone method due to the collecting-jar design. However, not all larvae emerged into the collecting jars, and we needed to dismantle the pyramids to find the larvae, which was time-consuming and likely led to larval mortality. We consistently observed desiccated larvae in the bottom of the pyramids, pointing to a humidity issue. However, when we attempted to raise the humidity within the pyramids, we had issues with mould growth that caused further mortality. Pyramid boxes are smaller and can be broken down and stored when not in use. They can also be set up in a temporary space or used in the field under ambient conditions. Therefore, despite our findings, this design could be useful for smaller experiments or where semi-permanent cages cannot be constructed, provided the issue of humidity regulation is addressed.
Emergence patterns
Daily larval emergence varied between locations and years (Fig. 5). Second-instar emergence usually began two to three days after we placed branches in flushing cages but took up to six days for some locations (Ontario, 2017), although this varied from year to year (Fig. 5). Once larvae emergence began, it peaked after 7–10 days, although we observed a peak in as little as five days (Fig. 5). Most of our larvae were collected 20 days after initiating flushing, so we would recommend this as a minimum length of time to ensure the majority of viable larvae were collected from each branch sample. Flushing continued until emergence ended or we obtained over 1000 second-instar larvae. As such, the length of flushing varied among populations. The density of overwintering second-instar larvae per branch affected the total number of insects obtained using either method. Given the variability observed among our different populations and between years, it is difficult to know beforehand how many branches would be needed or how long to flush branch bundles to obtain viable starting populations. High population densities provide more insects per branch than low-density populations, so prior knowledge of regional population dynamics would be useful to ensure an adequate number of branches are sampled.
Second-instar larval emergence by day for each population. Emergence presented as per cent of total emerged within a population. Day 1 indicates when branches were placed at flushing temperatures.
Seasonal weather differences experienced by spruce budworm larvae in 2016 and 2017 before their collection may account for the temporal differences observed in Ontario (Fig. 5), and similar variability has been documented before. Time of peak second-instar larval emergence varied among hosts (Lysyk Reference Lysyk1989) and between locations (Volney and Cerezke Reference Volney and Cerezke1992), with some sites showing the extended emergence (Volney and Cerezke Reference Volney and Cerezke1992) similar to our observations. Mortality of second-instar larvae can also be affected. For instance, warm fall temperatures and variable winter temperatures can lead to mortality during the second-instar stage (Han and Bauce Reference Han and Bauce1998; Marshall and Sinclair Reference Marshall and Sinclair2015). Substantial mortality occurred (> 50%) when second-instar larvae were exposed to several weeks of warm weather early at the onset of diapause (Han and Bauce Reference Han and Bauce1998). These factors have also been shown to affect emergence time (Saunders Reference Sanders1980). In addition, McMorran (Reference McMorran1973) showed that a laboratory colony of spruce budworm took an average of five days to emerge at 20 °C, but warm conditions before the onset of cool storage (“winter”) delayed second-instar larval emergence by several days. These and our own results suggest that a number of biotic and abiotic factors – including host, regional weather, and population differences – may influence second-instar emergence patterns and overwintering survival.
Rearing and quality control
The number of larvae established on diet and the overall survival varied among populations (Table 3), with mortality from emergence to eclosion ranging from 53% (Ontario 2018) to greater than 90% (Ontario in 2017, Québec, and New Brunswick; Table 3). To establish disease-free laboratory colonies, we eliminated diseased individuals and their progeny from our F0 generation. The combination of rearing mortality and cull of infected individuals led to cumulative mortality losses that amounted to 58–96% of flushed larvae (Table 3).
Demographics, disease incidence, and mortality in the F0 generation of six wild populations of spruce budworm used to establish laboratory colonies. Overall, infection rate and specific pathogen infection rate are based on the number of adults from single-pair matings (SPM)* that screened positive based on microscopic inspection of whole-body homogenate. Pre-screen mortality is the percentage of larvae that died before adult eclosion, and post-screen mortality is the percentage of insects lost before the F1 generation (i.e., after quality control). Number of clean adults and female sex ratio represent the individuals used to establish the F1 generation. NWT, Northwest Territories; AB, Alberta; ON, Ontario; NB, New Brunswick; QC, Québec; all locations are in Canada; NPV, nuclear polyhedrosis virus; EPV, entomopoxvirus.
*Single-pair matings used two to three adults, depending on adult availability.
Mortality differences across populations may be due to underlying fitness differences, variable natural environmental conditions before collection, and pathogen load (Bauer and Nordin Reference Bauer and Nordin1989). The Québec population was sampled in the fall, in contrast to the other populations that experienced winter temperatures before being sampled in the spring (Table 1). The Québec population showed the highest level of mortality (Table 3), but we cannot say whether this was due to the time of sampling or to other factors. Our wild populations also showed high variability in overall pathogen incidence (Table 3) and in specific pathogen types. The population from Alberta (Table 3) showed the highest overall incidence of disease (45.1%), whereas the populations from the Northwest Territories and Ontario (2018) had the lowest (10%; Table 3). Microsporidia and nuclear polyhedrosis virus were found in nearly all populations, whereas entomopoxvirus was detected only in the Ontario populations in 2017 (Table 3). We expected that disease incidence would vary between populations, but the regional variation we show here has been infrequently documented (but see van Frankenhuyzen et al. Reference van Frankenhuyzen, Nystrom and Tabashnik1995). Disease incidence can vary temporally throughout the outbreak cycle of spruce budworm (Eveleigh and Johns Reference Eveleigh and Johns2014), although the link between regional population dynamics and pathogen load is not clear; as a result, it is difficult to predict pathogen load in advance. Therefore, from the perspective of colony establishment, it is best to assume that substantial effort is needed to eliminate pathogens from new collections and to establish disease-free colonies.
Pathogens pose significant risks to laboratory colonies. Accidental introduction of pathogens into laboratory stocks can lead to colony collapse. Significant effort would then be required to recover disease-free stocks. One such collapse occurred in the Insect Production and Quarantine Laboratory in 2000, following the introduction of a microsporidian infection (van Frankenhuyzen et al. Reference van Frankenhuyzen, Ebling, McCron, Ladd, Gauthier and Vossbrinck2004). We assumed that our wild populations would harbour a range of pathogens, and we were therefore particularly cautious about introducing these wild populations to our rearing facility. In our wild populations, we detected Nosema fumiferana (Thomson) (Microsporida) (G. Kyei-Poku, personal communication), a species commonly found within spruce budworm populations (Bauer and Nordin Reference Bauer and Nordin1988 and references therein). Microsporidia, such as Nosema, can impact budworm fitness, are transmitted transovarially, and have a high degree of infectivity (Bauer and Nordin Reference Bauer and Nordin1989). However, the microsporidian we detected was a different species than the microsporidian that caused the previous colony collapse (van Frankenhuyzen et al. Reference van Frankenhuyzen, Ebling, McCron, Ladd, Gauthier and Vossbrinck2004). We also detected both nuclear polyhedrosis virus and entomopoxvirus within our F0 populations (Table 3). Both of these viruses occur naturally in the environment. They are ingested by insects during feeding, and infected larvae become active carriers (Bird Reference Bird1969; Whitlock Reference Whitlock1974; Podgwaite et al. Reference Podgwaite, Shields, Zerillo and Bruen1979). Visible symptoms of nuclear polyhedrosis virus include an unnatural brown or black colouration and fluid-filled sac-like appearance (Whitlock Reference Whitlock1974). Entomopoxvirus interrupts the hormone cascade that initiates metamorphosis, causing an oversized larva that cannot pupate and eventually dies (Palli et al. Reference Palli, Ladd, Tomkins, Shu, Ramaswamy and Tanaka2000). We detected low levels of cytoplasmic polyhedrosis virus in our F0 insects (data not shown). Cytoplasmic polyhedrosis virus is a latent virus that is common both in the wild and in captive insect colonies. Its presence does not usually impact fitness, unless it is aggravated by stress factors such as overcrowding (Mori and Metcalf Reference Mori and Metcalf2010).
We used three strategies to increase the survival of our emerged second-instar spruce budworms and to minimise the likelihood of disease transmission within these new laboratory stocks. First, we reared larvae individually to minimise horizontal disease transmission and to reduce competition for food (i.e., decreased stress will minimise cytoplasmic polyhedrosis virus expression). Second, we included fumagillin in the artificial diet, an antimicrobial used to manage microsporidial disease in spruce budworm, other Lepidoptera, and honeybees (Lewis and Lynch Reference Lewis and Lynch1970; van Frankenhuyzen et al. Reference van Frankenhuyzen, Ebling, McCron, Ladd, Gauthier and Vossbrinck2004). Third, we used single-pair or small-group matings, rather than mass matings, to reduce the risk of horizontal disease transmission to the F1 generation. Mass matings are more efficient in production scenarios (Ebling and Dedes Reference Ebling and Dedes2015) but are not appropriate for the initial establishment of wild colonies. Individual matings, although more time-consuming, allowed us to screen adults for pathogens. The disease state of each adult determined which progeny we retained for the F1 generation; we discarded progeny with a parent that tested positive for a pathogen unless only cytoplasmic polyhedrosis virus was present in small amounts. Although time-consuming, this method was effective in controlling the spread of microsporidia and viruses (van Frankenhuyzen et al. Reference van Frankenhuyzen, Ebling, McCron, Ladd, Gauthier and Vossbrinck2004), especially when we combined it with an antimicrobial agent in the diet (Table 3). Our data suggest these efforts were successful at lowering the incidence of infection in the F1 generation. We did not detect microsporia in our F1 populations, with the exception of the population from Alberta, in which we detected the pathogen in 7.4% of screened adults. Although fumagillin and single matings were effective at reducing disease effects on the F0 generation, continued use of the antimicrobial agent caused physical deformities in the F1 generation (Fig. 6). We did not observe these deformities in F1 larvae reared on a fumagillin-free diet (Fig. 6; data not shown). Studies have demonstrated that multigenerational use of the antimicrobial agent can cause a reduction in larval survival, pupal weights, and fecundity (van Frankenhuyzen et al. Reference van Frankenhuyzen, Ebling, McCron, Ladd, Gauthier and Vossbrinck2004), and they caution against the use of fumagillin for more than one generation, despite its effectiveness.
Pupal deformities associated with fumagillin use within the diet. A, Normal pupae; B–C, pupae displaying developmental abnormalities.
Our precautions regarding the use of fumagillin may be somewhat moot, as the sole producer of fumagillin ceased production in 2019. Other antimicrobials are less effective for managing latent microsporidial infections (van Frankenhuyzen et al. Reference van Frankenhuyzen, Ebling, McCron, Ladd, Gauthier and Vossbrinck2004). Because of this, the loss of this product will likely hamper future efforts to establish spruce budworm and other insects as laboratory colonies. Our evidence suggests that if other antimicrobials are developed, the effects on F0, F1, and subsequent generations should be investigated during the colony-development process.
Genetic considerations
Laboratory conditions are drastically different from field environments and can have unintended effects on the newly established colonies. Through the process of selection and genetic drift, laboratory colonies can change relative to their wild ancestors, both genetically and functionally (Caprio Reference Caprio2009; Berthier et al. Reference Berthier, Chapuis, Simpson, Ferenz, Kane and Kang2010). Genetic drift is a random change in gene allele frequencies over time, whereas selection occurs because of adaptation to the laboratory environment. Genetic drift will affect the entire genome equally; maintaining a large population size is therefore the best means to mitigate its effects (Caprio Reference Caprio2009). Our goal was to establish new spruce budworm colonies that reflect the genetic and phenotypic variation of this insect in Canada. We initiated colonies from at least 33 individuals and maintained a female sex ratio of 1:1 (range 0.6–1.0), with most populations established with 100 or more mated pairs (Table 3). For some colonies, we added F0 individuals in a subsequent year (e.g., Glfc:IPQL:CfumTON01; Table 3), but we were unable to do so for others, particularly those from remote locations (e.g., Glfc:IPQL:CfumINT01; Table 3). We had sought to establish the F1 generation from at least 100 disease-free single-mated pairs to minimise the effects of drift; however, mortality and disease in our first generation limited our founding population. Similar population sizes were used to re-establish the colony at the Insect Production and Quarantine Laboratory following a genetic bottleneck caused by disease (van Frankenhuyzen et al. Reference van Frankenhuyzen, Ebling, McCron, Ladd, Gauthier and Vossbrinck2004; Roe et al. Reference Roe, Demidovich and Dedes2018), with no observed negative consequences. Furthermore, our founding laboratory populations likely capture the majority of the genetic diversity within the source populations (Berthier et al. Reference Berthier, Chapuis, Simpson, Ferenz, Kane and Kang2010), based on the previously published estimates of spruce budworm allelic richness and heterozygosity (James et al. Reference James, Cooke, Brunet, Lumley, Sperling and Fortin2015; Larroque et al. Reference Larroque, Legault, Johns, Lumley, Cusson and Renaut2019). We used a minimum of 100–200 individuals of each sex to generate the next generations of each colony, which should help to maintain the founding genetic diversity for at least 100 generations (Berthier et al. Reference Berthier, Chapuis, Simpson, Ferenz, Kane and Kang2010). Successful establishment from founding populations of more than 100 individuals is common in biological invasions and biological control studies, whereas founding populations of fewer than 10 are more likely to fail. Because we founded most of our populations from more than 100 adults, and all colonies were initiated with more than 10 individuals, we are confident that these laboratory-reared colonies are representative and sustainable.
Selection is the other primary cause of genetic change in laboratory colonies. Unlike genetic drift, selection acts on specific genes and loci within the genome. Selection occurs naturally as insects adapt to laboratory conditions, and the degree of change depends on the strength of selection (Berthier et al. Reference Berthier, Chapuis, Simpson, Ferenz, Kane and Kang2010). Selection is more difficult to manage, given the selective pressures inherent in the transition from natural to artificial conditions. Often new laboratory colonies collapse after several generations because of high selective pressures (Godinho et al. Reference Godinho, Cruz, Charlery de la Masselière, Teodoro-Paulo, Eira and Fragata2020). Outbreeding is a common means to create robust colonies and to minimise selection by maximising standing genetic variation through the introduction of new genetic material from other populations (Godinho et al. Reference Godinho, Cruz, Charlery de la Masselière, Teodoro-Paulo, Eira and Fragata2020). However, outbreeding is not compatible with our goal to study local adaptation. Alternatively, infusing the laboratory colonies with new field-caught specimens would also help counteract selective pressures on our new colonies. However, this process is time and resource intensive. Quality control assessments of performance traits can help to assess whether laboratory insects are deviating from the norms defined in wild populations and could help identify whether laboratory colonies would need genetic infusion to improve performance. Establishing baseline values for insect performance (i.e., fecundity, pupal size, and mortality rates) can point to selective change that is occurring within a laboratory population. Preserving a genetic record of the original wild stock can provide a valuable comparison for assessing the degree of change within laboratory populations (Caprio Reference Caprio2009). To this end, we kept frozen tissue vouchers of insects from each population to use as a genetic record of the founding population. Going forward, we will use fitness and genetic quality assessments to document potential change in our new spruce budworm colonies over time.
Conclusion
Our goal was to establish new spruce budworm colonies from across the species’ range in Canada. Although rearing protocols for maintaining existing spruce budworm colonies are well established, we found that the processes needed to establish new laboratory colonies were poorly described in the literature or discussed in isolation. Here, we describe the complete process of establishing new colonies of the insect – from field collection to the production of disease-free laboratory stocks. We found that paper cones were more effective for flushing large numbers of insects from field-collected branches. Disease and artificial laboratory conditions often combine to result in high mortality during the initial stages of rearing, thereby presenting significant challenges to colony establishment. We used single-pair matings and antimicrobials to manage pathogens, which led to substantially lower incidence of disease in the F1 generation. Despite these measures, the initial F0 generation experienced high mortality. Given the rate of loss, we recommend starting with large numbers of F0 insects and minimising larval mortality to ensure adequate numbers of adults for the founding generation. A large founding population will adequately capture the genetic diversity within wild populations, help minimise drift, and ensure colony longevity.
Acknowledgements
The authors express their gratitude to all those who helped them develop the new spruce budworm colonies. For collection of wild populations, these individuals include M. Callaghan (Northwest Territories Government), C. Whitehouse and R. Hermanutz (Alberta Agriculture and Forestry), R. Brett and G. Pohl (Canadian Forest Service – Northern Forestry Centre, Edmonton, Alberta, Canada), M. Grey and G. Brand (Canadian Forest Service – Great Lakes Forestry Centre, Sault Ste. Marie, Ontario, Canada), L. De Grandpré and staff (Canadian Forest Service – Laurentian Forestry Centre, Québec, Québec, Canada), and R. Johns and staff (Canadian Forest Service – Atlantic Forestry Centre, Fredericton, New Brunswick, Canada). For sharing their knowledge of spruce budworm flushing methods, the authors thank R. Scharbach and T. Ladd (Canadian Forest Service – Great Lakes Forestry Centre). They also thank R. Fournier (Canadian Forest Service S – Great Lakes Forestry Centre) for developing the map of spruce budworm defoliation, G. Kyei-Poku (Canadian Forest Service – Great Lakes Forestry Centre) for confirming the microsporidial species within the wild populations, B. Gribelli (Canadian Forest Service – Great Lakes Forestry Centre) for keeping the insect chambers running smoothly, and J. Dedes, K. van Frankenhuyzen, and staff at the Insect Production and Quarantine Laboratory, Great Lakes Forestry Centre for their helpful advice, guidance, and support during this process. This work was funded by the Spray Efficacy Research Group International (SERG-I) and its provincial partners (Grant no. 2016/06-2016-114).
