Using a portable vacuum cleaner coupled to a simple acrylic tube custom made with a soft net on one end (Figure 1B), we collected large numbers of flies from a vineyard dump site (Figure 1C). After collection, flies were transferred directly to bottles containing standard cornmeal-agar medium (Figure 1D). Around 1000 females were separated and individually distributed to vials with standard food. These females were transferred twice to new vials, laying eggs during 3 days in each vial.

To perform nucleic acid extraction, after the second round of egg laying, each vial with fertile progeny (checking the presence of F1 larvae in the food) was numbered and the respective female was anesthetized with CO₂. A 96-well plate was previously prepared for DNA extraction, cooled over dry ice to facilitate the placing of the anesthetized females in the respective wells.

Nucleic acid extraction was performed according to (http://www.drosdel.org.uk/molecular_methods.php) with minor modifications. Briefly, each biological sample was homogenized with metallic beads and detergent for cell lysis. After removing the cellular waste by centrifugation, the supernatant was transferred to a new 96-well plate and nucleic acids were separated and precipitated with KCl and isopropanol. Samples were further washed in EtOH 70% and afterwards ressuspended in milliQ H₂O. All steps were alternated with centrifugation steps. DNA or RNA were isolated after incubation with either RNase or Dnase, respectively.

After DNA extraction, a multiplex PCR reaction was performed in 96-well plates using 1 μL of diluted DNA, GoTaq DNA polymerase (Promega) in a 20 μL total reaction volume per well, using the primers Slif (Fwd–5′GTTAGCGCCTATTAGCACAT; Rev–5′CGGGACAACTCAGTCTGTAA) to distinguish between D. melanogaster and D. simulans, and wsp (Fwd8 1-5′-TGGTCCAATAAGTGATGAAGAAAC-3′; Rev691-5′-AAAAATTAAACGCTACTCCA-3′), to diagnose for the presence or absence of Wolbachia. The PCR program was used as follows: 95°C for 10 min; 30 cycles at 95°C for 30 s (denaturation), 60°C for 1 min (annealing), 72°C for 1 min (elongation) and a final extension step at 72°C for 10 min. PCR amplification products were visualized after electrophoresis in agarose gel (1.5% in TAE supplied with 0.5% RedSafe).

After analysis, the F1 vials of each tested-isofemale line could be separated into four groups, D. simulans or D. melanogaster, and Wolbachia positive or negative.

To create a Wolbachia-negative population from the founded outbred infected population, an egg lay with controlled density was allowed in standard food containing 0.05 mg/ml tetracycline hydrochloride (Sigma). Throughout 3 generations, the full life cycle was carried out in food containing tetracycline at the same concentration, after which the population returned to regular maintenance in standard food for two generations for mitochondrial recovery. Finally, we standardized the microbiota by transferring the gut microbiome from the original population (Wolbachia-positive), as described in Faria et al. (2016). In the case of isofemale lines, the same procedure can be applied and adjusted to tubes.

From 160 isofemale lines of each group previously diagnosed, 10 F1-flies, 3–5 virgin females and 3–5 males were separated. Groups of males and virgin females were placed simultaneously in population acrylic boxes (50 × 30 × 25 cm), thus minimizing sib-mating. Around 1,500 individuals founded each population.

Populations were kept on a 3-week non-overlapping generation cycle. Treatments were always performed 3–5 days after eclosion and reproduction occurred 5–7 days after treatment. Reproduction was performed in 10 plastic cups (5 per day) with standard food. Egg density was limited to 400 per cup, a density determined experimentally to enable optimal larval development and population effective numbers. Flies were maintained under constant temperature (25°C), humidity (60–70%) and light-darkness cycle (12:12 h), and fed with standard cornmeal-agar medium.

Each population was kept in laboratory cages with high census (between 1,500 and 2,000 individuals). Census above 2,000 flies will lead to excess moisture inside the boxes that compromises egglaying, mobility and viability of flies and promotes bacterial and fungal proliferation.

Average expected heterozygosity in Initial population (2007) and outbred population adapted to the laboratorial conditions (2010), were estimated using 103 SNP markers described in Teotónio et al. (2009) and were analyzed as described in (Martins et al., 2013; Supplementary Information from Martins et al., 2014). We used samples of 52 individuals from initial founding females and 48 individuals in 2010.

Before the initiation of experimental evolution experiments, populations were maintained under the laboratory previously described conditions for a minimum of 15 generations and then serially expanded for two generations to allow the establishment of the new replicate populations. In our case, all lines of all treatments were derived from the same base population (from 1 to 6 in first generations and from 6 to 36 in the second). It enables that all populations under different experimental treatments will start with the same genetic pool, even losing some initial variability due to adaptation to the laboratory or to an unequal representation of founder lines. The egg laying for the foundation should be randomly distributed across the replicates to avoid any selection for fertility. In each generation, pools of 200–300 flies of each replicate were frozen in liquid nitrogen and kept at −80°C.

Isofemale lines were established using the vials of each original line used to extract the virgins and males which have founded the outbred population. In this case the lines were maintained, as is customary, with overlapping generations.

Starting from the isofemale lines, one can also establish isogenic lines. To do so, each isofemale line can be taken through 20 generations of full-sib mating as done by Mackay et al. (2012). This procedure should purge deleterious alleles and provide, at a reasonable frequency, fertile and viable genetically-homogeneous lines.

We introgressed the white mutant allele (w¹¹¹⁸) into the outbred population. Using 80–100 single female crosses in the first two generations, we replaced all second and third chromosomes from the w¹¹¹⁸ stock by “wild” chromosomes of the outbred population. In each odd generation (F1, F3, F5, F7, F9, F11) recombination in females reduces the contribution of the w¹¹¹⁸ stock. From these crosses, 2 white-eyed males were used to establish at least 100 single female crosses with virgins from the outbred stock (all even generation F2, F4, F6, F8, and F10). In the F12 generation, 3–5 virgin females and 3–5 males from 140 F11 single-female crosses were released into a population cage to establish the white introgressed population.

Crossing scheme:

Number of individuals used in each single-female cross throughout the introgression procedure.

Generic methods to collect Drosophila species have been described in Markow and O'Grady (Markow, 2006). For the specific collection of Drosophila melanogaster, our own experience favors the choice of a vineyard as the collection site, given the advantages that collecting from large populations provides (particularly during harvest, which corresponds roughly to the period from August to October in the northern hemisphere) (Figure 1). Using a hand vacuum cleaner adapted to this purpose (see M&M), in 2007 and 2013, we collected in 1h around 5000 females from the José Maria da Fonseca winery in Southern Portugal (Azeitão, Portugal GPS: 38° 31′ 04.91″ N9° 00′ 56.24″).

Back to the lab, we followed systematically the steps schematically represented in Figure 2 in order to maximize the collected resources. First, single females were separated in vials to lay eggs and ensure the next generation. Then, the progenitor females were sacrificed and used as starting material for individual nucleic acids extraction (both DNA and RNA) in 96-well plates.

Guided by Alberto Civetta (Minuk and Civetta, 2011), we have scanned the genomes of D. melanogaster and D. simulans for large indels that would be diagnostic of species through a simple PCR followed by electrophoresis. We chose Slif (CG11128) for which we designed a primer pair (see M&M) that amplifies fragments of 939 bp in D. melanogaster and 1058 bp in D. simulans. To detect Wolbachia, we used primers designed for wsp (Wolbachia surface protein) by Zhou et al. (1998) (see M&M), which produce two differently sized fragments of 610 and 590 bp, for D. melanogaster and D. simulans, respectively. The Wolbachia wsp gene amplification product serves therefore two purposes: the characterization of the individual's infection status and a second (indirect) confirmation of the Drosophila species itself, conveyed by the different amplicon sizes generated by respective Wolbachia strains.

Figure 3 shows how this combination of primers reveals simultaneously, in a multiplex PCR (see M&M), the status of Wolbachia infection and which of the cryptic species it is. The PCR program used permits the simultaneous amplification of both fragments without any primer incompatibility or confounding effects from nonspecific bands. We have performed this method at high throughput rate using 96-well plates.

To validate the species diagnosis, we performed in parallel a blind test running our method and performing the classical male genitalia classification based on morphological analysis. For this we took 50 males from one collection and mounted their genitalia (Figure 4) as described in Ashburner (Ashburner et al., 2011) and used the carcass to prepare genomic DNA for each individual. We found a 100% match (50 in 50) fit between the two classification methods, leading to the conclusion that the method we developed is at least as good as the more time-consuming (and more subjective) classical method.

As an illustration of the proportions found in our specific case, in the 2013 collection we genotyped 576 (96 × 6) fertile females (progenitors of isofemale lines), being 341 (D.mel/Wol+), 189 (D.mel/Wol−), 36 (D.sim/Wol+), 1 (D.sim/Wol+), and 9 failed extractions/amplifications.

From the collection described above, we established an outbred population from 160 fertilized Wolbachia-positive females. We used 3–5 virgin females and 3–5 males from the F1 of each previously screened line. In parallel, we started 160 isofemale lines, kept in similar maintenance conditions but under an inbreeding regime, in vials with uncontrolled census and overlapping generations. Further, after 3 years we re-derived an outbred population without Wolbachia through tetracycline treatment (see M&M). After the full procedure, the new population (Wolbachia-free) was ready to be expanded and tested.

We performed extensive analyses for quality-control purposes of these established populations. Specifically, we quantified heterozygosity (H) and Fst for a subset of the starting foundation lines (using the extracted and stored DNA as described above) and in the two established outbred populations. We used the genomic DNA extracted from the founding females in 2007 to genotype 48 individuals and estimated an Expected Heterozygosities (H_exp) of 0.344 (±0.016 SE). In 2010 we repeated the procedure on the established outbred population to find a H_exp of 0.320 (±0.017 SE) (Martins et al., 2014). We estimated the Fst at 0.038.

After the establishment of laboratory-controlled outbred populations, namely of Drosophila melanogaster infected with Wolbachia, we waited several generations for adaptation to the lab and consecutive stabilization of genetic pool. After that, we sequentially expanded the population in order to generate several identical replicas (see M&M). In our case, we used those populations to follow their response and genetic configuration upon different pathogenic challenges and different infection routes (Martins et al., 2013, 2014). Additionally, we also approach the eventual costs of adaptation and the influence of the increase of immunocompetence on Wolbachia population (Faria et al., 2015) and the evolutionary significance of its presence (Faria et al., 2016).

In each generation of experimental evolution, 200–400 flies (males and females) were frozen and posteriorly used for different analyses, such as PoolSeq and genotyping (Martins et al., 2014; Faria et al., 2016).

After the foundation of outbred populations, the introgression of specific alleles in the populations can be performed. In the case of visible markers this procedure is rather simple, though laborious, as exemplified by the scheme below used to introgress the white mutant allele (w¹¹¹⁸) into the outbred population. Using over 100 single female crosses per generation, we recombined the white mutant allele into “wild” chromossomes 6 times. Also, all other chromosomes were outbred as they were replaced in full (using balancer chromosomes) from the initial generations of the crossing scheme (see M&M). With this number of recombination rounds the estimated proportion of the white allele-carrying X chromossome in the final population is theoretically inferior to 2% (2^∧6 = 1/64). Except for the fragment of the X in linkage with the white locus, every other fragment from the w¹¹¹⁸ stock was randomized and should have virtually no impact on the phenotypes observed at the population level. Effectively, we have generated an outbred population genetically indistinguishable from the wild-derived initial outbred population except for the fact it carries the w¹¹¹⁸ allele.

Additionally, we have started a systematic introgression of transgenic lines into the outbred background using the same crossing scheme.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.